[new comprehensive biochemistry] calcium - a matter of life or death volume 41 || ryanodine receptor...

J. Krebs and M. Michalak (Editors)

Calcium: A Matter of Life or Death

� 2007 Elsevier B.V. All rights reserved.

ISSN: 0167-7306/DOI: 10.1016/S0167-7306(06)41012-7

CHAPTER 12

Ryanodine receptor structure, functionand pathophysiology

Spyros Zissimopoulos and F. Anthony LaiDepartment of Cardiology, Wales Heart Research Institute, School of Medicine, Cardiff

University, Cardiff CF14 4XN, UK, Tel.: +44 29 207 42338; Fax: +44 29 207 43500;E-mail: [email protected], [email protected]

Abstract

The ryanodine receptor (RyR) is an intracellular calcium release channel located on thesarco(endo)plasmic reticulum of muscle and non-muscle cells. The functional channel is

composed of four identical subunits of approximately 560kDa, which combine to form ahigh-conductance cation-permeable protein pore. There are three mammalian RyR iso-forms that have a wide tissue expression. Their highest levels are in striated muscles where

they mediate the release of stored Ca2+ leading to a rise in intracellular Ca2+ concentra-tion and muscle contraction. Channel activity is regulated by Ca2+, Mg2+, ATP and post-translational modifications, i.e. oxidation/reduction and phosphorylation. In addition, the

RyR is regulated by intramolecular protein–protein interactions, as well as by interactingwith numerous accessory proteins including the dihydropyridine receptor (DHPR),FK506-binding protein (FKBP), calmodulin (CaM), sorcin and calsequestrin (CSQ).Inherited or acquired defective channel regulation results in abnormal Ca2+ handling

and leads to neuromuscular disorders and arrhythmogenic cardiac disease.

Keywords: calcium release channel, calmodulin, calsequestrin, DHPR, E–C coupling,

FK506BP, heart failure, neuromuscular disorder, phosphorylation, redox status, ryano-dine, ryanodine receptor, RyR accessory proteins, RyR pathophysiology, sarco(endo)-plasmic reticulum, sorcin

1. Introduction

The ryanodine receptor (RyR) is a family of Ca2+ channels located in the mem-branes of internal Ca2+ storage organelles. It is closely related to the other family ofintracellular Ca2+ channels, the inositol trisphosphate receptor (IP3R), with whichthey provide a regulated pathway for the release of stored Ca2+ during Ca2+

signalling processes such as muscle contraction and fertilization (reviewed in [1]).The RyR and IP3R proteins share structural similarities, high homology especially attheir carboxyl-terminal sequences that form the channel pore (approximately 40%homology) and functional similarities including Ca2+ and ATP regulation.

RyRs were initially observed in skeletal muscle in the early 1970s, where they werevisualized in electron micrographs as large electron-dense masses situated along the

face of the sarcoplasmic reticulum (SR), spanning the junctional gap between SR andthe plasma membrane, and they were therefore termed junctional foot proteins(reviewed in [2]). The RyR gained its present name in the late 1980s after it wasfound to be the protein that binds ryanodine, a plant alkaloid that enabled purifica-tion and molecular characterization of the protein.

Ryanodine is a natural product found in members of the genus Ryania, whichgrow as shrubs or slender trees in several tropical locations in Central and SouthAmerica, with potent paralytic actions on skeletal and cardiac muscles (reviewed in[3]). Crude ryanodine extracts were originally used to prepare poison arrowheads,and in the 1940s, powdered Ryania wood was marketed as an insecticide. A com-pound purified from stem wood was found to have 700 times the insecticidal potencyof the starting material and was designated ryanodine. Ryanodine binds to the RyRwith high affinity and specificity, preferably in its open conformation, and thus,ryanodine binding is used as an index of channel activation. The RyR is the onlycellular target of ryanodine reported to date.

The RyR was purified from skeletal and cardiac muscle using [3H]–ryanodine as aselective marker and found to exist in a tetrameric form that sediments as a very large(30 S) complex by sucrose density gradient centrifugation [4–11]. The purified pro-tein, incorporated into an artificial planar lipid bilayer, functions as a Ca2+ channel,with characteristics identical to the Ca2+ release channel observed upon incorpora-tion of the crude SR fraction [6,8,10,12,13].

2. RyR isoforms and distribution

2.1. Isoforms

Biochemical, molecular or pharmacological evidence for the presence of RyRs hasbeen found in vertebrates (mammals, birds, amphibians, reptiles and fish) as well asinvertebrates including crustaceans, insects and nematodes (reviewed in [14,15]).

In mammals, three RyR isoforms encoded by distinct genes have been cloned andfully sequenced, RyR1 in skeletal muscle [16,17], RyR2 in heart [18,19] and RyR3 inbrain [20], revealing subunit composition of approximately 5000 amino acids andmolecular weight of approximately 560kDa (Table 1). The currently accepted terminol-ogy is based on the abundance and timing of purification of the RyRs from varioustissues. Thus, RyR1 is also known as the skeletal type, RyR2 is also known as thecardiac type, whereas RyR3 was initially but misleadingly termed the brain type. ClonedRyRs have been heterologously expressed in various mammalian cell lines includingCHO, HEK293 and COS-1 [16,21–26]. Expressed proteins were indistinguishable fromthe native channels in immunoreactivity, molecular size, sucrose density gradient sedi-mentation and ryanodine-binding properties. Furthermore, recombinant channels hadconductance, kinetics of opening, current–voltage relationship, cation permeability andmodulation by physiological and pharmacological ligands similar to the native channels.

The overall percentage identity of the three mammalian RyRs is approximately70% (Table 1), with the highest sequence identity present at the carboxyl-terminal

288 S. Zissimopoulos and F. A. Lai

Table 1

Homology between ryanodine receptor isoforms

RyR1human

RyR1pig

RyR1rabbit

RyR1fish

RyR1frog

RyR2rabbit

RyR2human

RyR2mouse

RyR3human

RyR3rabbit

RyR3mink

RyR3chicken

RyR3frog

RyR

seaurchin

RyR

Drosophila

RyR

Caenorhabditis

elegans

Length(aa)

GenBankaccession

number

RyR1 human 100 5032 J05200

RyR1 piga 96 100 5035 M91452

RyR1 rabbit 96 96 100 5037 X15750

RyR1 fish 74 74 74 100 5081 U97329

RyR1 frog 78 78 78 78 100 5037 D21070

RyR2 rabbitb 65 65 65 64 66 100 4969 M59743

RyR2 human 65 65 65 64 66 98 100 4967 X98330

RyR2mouse 64 64 64 63 66 97 97 100 4968 AF295105

RyR3 humanc 64 64 64 63 66 67 66 67 100 4866 AB001025

RyR3 rabbit 64 64 65 63 66 67 67 67 95 100 4872 X68650

RyR3mink 64 65 65 64 67 67 67 67 95 95 100 4859 Y07749

RyR3 chicken 64 64 64 64 66 67 67 67 86 86 86 100 4869 X95267

RyR3 frog 65 64 65 64 68 68 68 67 85 85 85 85 100 4868 D21071

RyR sea urchin 44 44 44 43 44 46 45 45 34d 33d 33d 34d 34d 100 5317 AB051576

RyR Drosophila 44 44 44 42 43 45 45 45 34d 33d 34d 35d 35d 44 100 5126 D17389

RyR Caenorhabditis elegans 39 39 39 38 38 41 41 41 34d 35d 35d 32d 33d 40 45 100 5071 D45899

Table illustrating the percentage sequence identity of the various optimally aligned ryanodine receptors (RyRs). The sequence alignments between RyR pairs

were produced with the ‘pairwise BLAST’ software available at the NCBI website (http://www.ncbi.nlm.nih.gov).aA second pig RyR1 cDNA sequence has also been published (X62880); at the protein level, the second sequence presents a deletion of one residue and five

additional mismatches.bA second rabbit RyR2 cDNA sequence has also been published (U50465); at the protein level, the second sequence presents a deletion of one residue and six

additional mismatches.cA second human RyR3 cDNA sequence has also been published (AJ001515); at the protein level, the second sequence is four residues longer due to two

alternative splicing events: the first involves replacement of 26 amino acids by a distinct 25-residue fragment and the second involves insertion of five amino

acids and there are also 18 additional mismatches.dVertebrate RyR3 isoforms do not align with sea urchin, Drosophila and Caenorhabditis elegans RyRs over the entire sequence but only over fragments of low

identity; the figure given is the average one.

end. Lower overall percentage identity is due to three regions of marked divergence(D1, D2 and D3) [27]. With reference to the RyR1 sequence, D1 spans amino acids4254–4631, D2 spans amino acids 1342–1403 and D3 lies between residues 1872 and1923. D1 is the largest and most variable region, whereas D2 is completely absent inRyR3.

Non-mammalian vertebrates (birds, amphibians, reptiles and fish) also expressthree subtypes, initially called the a-type, b-type and the cardiac type [28–33]. On thebasis of sequence comparisons with the mammalian isoforms, the a- and b-RyRs areidentified as the mammalian RyR1 and RyR3, respectively [34,35]. Only one type ofRyR has been detected in invertebrates including lobster [36], fly [37] and nematode[38]. Full-length cDNA sequences are known for sea urchin (Hemicentrotus pulcher-rimus), fly (Drosophila melanogaster) and nematode (Caenorhabditis elegans), whichshow 35–45% similarity with the vertebrate RyRs, the highest similarity with theRyR2mammalian isoform (Table 1). Phylogenetic analysis suggests that the RyR2isoform diverged from a single ancestral gene before RyR1 and RyR3 isoforms toform a distinct branch of the RyR family tree [39]. The RyR2 is also the subtype withthe highest conservation across species (Table 1).

Alternative splicing that could introduce functional complexity in the RyR familyhas been described [18,39–44]. The resulting RyR proteins may have unique func-tional roles as some splice variants are expressed in a tissue- and developmentalstage-specific manner. Indeed, a dominant negative RyR3 splice variant, which lacksresidues 4406–4434 encompassing a putative transmembrane (TM) domain, has beendescribed in smooth muscle tissues but not in skeletal muscle, heart or brain [44].Heterologous expression of the splice variant did not result in a functional channel,whereas co-expression with wild-type RyR3 resulted in heteromeric channels withsuppressed activity compared to wild-type homotetramers. In addition, aberrantsplicing of RyR1mRNA has been implicated in impaired Ca2+ homeostasis inmyotonic dystrophy type 1 (DM1) muscle [45]. It was shown that the foetal RyR1variant, which lacks residues 3481–3485, was significantly increased in DM1 skeletalmuscle and exhibited reduced activity compared to the wild-type protein. Additionalcomplexity may arise from the formation of mixed heterotetrameric RyR channels.An immunoprecipitation study of recombinant, heterologously co-expressed RyRsprovided evidence that RyR2 could form heterotetramers with RyR1 and RyR3 butnot RyR1 with RyR3 [46]. However, the presence of native tissue RyR heterotetra-mers has not so far been reported.

2.2. Tissue and cellular distribution

Biochemical, molecular or pharmacological evidence indicate that RyRs have awidespread expression including skeletal, cardiac and smooth muscles, neurons ofboth the central and peripheral nervous systems, liver, lung, kidney, testis, ovary,endothelial, epithelial, pancreatic and adrenal chromaffin cells, osteocytes, neutro-phils and macrophages (reviewed in [14,15]). The RyR is primarily located on the SRof muscle cells and the endoplasmic reticulum of non-muscle cells. It has also been


reported that the RyR is found in mitochondria, where it mediates mitochondrialCa2+ uptake [47], and in secretory vesicles of a pancreatic b-cell line [48].

The highest RyR levels exist in striated muscle, RyR1 in skeletal and RyR2 incardiac muscle, whereas all three isoforms are expressed in brain and several smoothmuscles [16,18–20,28,30,31,33,49–56]. In mammals, the RyR1 isoform is the predo-minant form expressed in skeletal muscle and in some parts of the brain, mostprominently in Purkinje cells of the cerebellum. The RyR2 isoform is the predomi-nant form in cardiac muscle and is also the most widely and abundantly distributedisoform in the brain. RyR3 is expressed in a wide range of cells including the brain,skeletal and smooth muscles and epithelial cells, but at low levels. In contrast tomammalian skeletal muscle, amphibian and avian skeletal muscles express bothRyR1 and RyR3 (the a- and b-types, respectively) at approximately equal amounts.It is now appreciated that the myocardium is the source with the purest RyRcomposition (RyR2), whereas mammalian skeletal muscle is the richest source ofRyR1 with a minor contamination with RyR3. The highest levels of RyR3 inmammals have been reported in diaphragm muscle, but they still represent <5% ofthe overall RyR population [57,58].

2.3. Gene knock-out studies

The physiological role of the RyR has been addressed by gene knock-out studies.Mice deficient in RyR1 died at birth with gross abnormalities of the skeletal muscleand had abolished contractility following electrical stimulation [59]. Similarly, micedeficient in RyR2 died at the embryonic stage due to severe cardiac defects andimpaired Ca2+ homeostasis although there was apparently normal excitation–con-traction (E–C) coupling [60]. Both RyR1- and RyR2-deficient mice presentedmarked degeneration of skeletal and cardiac muscle, respectively, indicating theimportance of these proteins in normal muscle maturation. On the other hand,embryonic lethality did not allow investigation of the role that RyR1/2 could playin other tissues. In contrast to RyR1/2 knock-out animals, mice deficient in RyR3were viable with no gross abnormalities and exhibited functional E–C coupling inadults [61,62]. However, skeletal muscle contraction was impaired during the firstweeks after birth suggesting a role for RyR3 in neonatal skeletal muscle. In addition,physiological and behavioural studies have revealed that the RyR3-deficient miceexhibit altered hippocampal synaptic plasticity and impairment of learning capabil-ities [63,64].

3. Channel properties

Ion conduction through the RyR, being an intracellular channel, cannot be mon-itored using standard whole-cell recording techniques, but it can be monitoredfollowing incorporation of the channel into artificial lipid bilayers (single channelrecordings). Although RyR is impermeable to anions, the channel is permeable to a

Ryanodine receptor structure, function and pathophysiology 291

wide range of inorganic divalent and monovalent cations as well as organic mono-valent cations (reviewed in [65]). The unitary channel conductance for the threemammalian RyR subtypes is similar and extremely high, approximately 10-foldhigher than the plasma membrane Ca2+ channel [66–70]. Despite channel conduc-tance being generally lower with divalent (e.g. 135–150 pS for Ca2+) than mono-valent cations (e.g. 720–800 pS for K+) as the charge carrier, divalent cations are sixto seven times more permeable than K+. Hence, the RyR selects rather poorlybetween cations, and it is certainly not as selective as the plasma membrane Ca2+

channel. However, Ca2+ is essentially the only ion released from the SR, because it isthe only cation with a significant concentration gradient across the SR membrane. Itappears that due to its localization, the RyR has evolved into a high-conduction porewith little need for ion discrimination.

The RyR1 is permeable to relatively large compounds including choline andxylose (radius approximately 2.9 A and approximately 3.4 A, respectively) suggestinga rather wide pore [66,71]. Molecular sieving experiments using organic monovalentcations of increasing size have indicated that the minimal width of the RyR2 pore isapproximately 3.4 A [72], in agreement with streaming potential measurements indi-cating that H2O (radius 1.5 A) and Cs+ (radius 2 A) must pass in a single file fashion[73]. Electrical distance measurements using trimethylammonium derivatives of vary-ing length indicated that the length of the voltage drop along the RyR2 channel poreis approximately 10.4 A [74], consistent with streaming potential measurementsshowing that three H2O molecules (approximately 9 A) occupy the single file regionof the pore [73]. Voltage-dependent block of ion translocation by organic compoundssuch as tetramethylammonium and triethylamine (radius 3.6 A) was found to occurat a site located 90% into the voltage drop from the cytosolic face of the channel[72,75]. These investigations indicate that the RyR pore is effectively a wide tunnelwith a short constriction of approximately 7 A in diameter near its luminal exit, and atotal length of approximately 10 A. Thus, the RyR provides a pathway for cationtranslocation across the SR membrane (45–50 A) through a short pore (approxi-mately 10 A) that is flanked by large cytosolic and/or luminal mouths or vestibules.

Several studies have indicated the presence of negatively charged residues on boththe cytosolic and the luminal ends of the conductance pathway of the RyRs. Addi-tion of carboxyl-neutralizing chemical modifiers to the luminal face of RyR2 sig-nificantly reduced channel conductance [76]. Large organic and inorganic cations,anaesthetics and peptides were shown to block ion translocation through RyR1 andRyR2 channels from the cytosolic side [75,77–79], whereas polycations such asneomycin and ruthenium red were capable of blocking the channel from both sides[80,81]. It has been proposed that the high density of negative charge at both mouthsof the RyR pore serves not only to deny access to anions but to increase the localcation concentration ensuring a high-delivery rate to the pore, whereas it could alsocontribute to the relative permeability of divalent over monovalent cations [82].

The tetrameric RyR channel comprises a single pore although sublevel conduc-tance states corresponding to one-fourth, one-half and three-fourth of the mainconductance level for the purified channel have been reported [66]. Four subconduc-tance states have also been detected following dissociation of the small accessory


FK506-binding protein 12 (FKBP12), which led to the proposal that the channelcontains four pores with each subunit contributing an individual ion-conductingpathway [83]. However, single pore channels can also exhibit subconductance statesbecause of uncoordinated monomers. Evidence for a single pore is provided by astudy that combined two approaches: modification of the RyR2 pore by metha-nethiosulfonate ethylammonium (MTSEA+) to reduce current amplitudes and theuse of different-sized current carriers [84]. MTSEA+ modification decreased thenumber of channel substates as the diameter of the current carrier increased, indicat-ing that the conduction pathway is comprised of a single central pore. Furthersupport for a single ion-conducting pathway comes from heterologous co-expressionof wild-type RyR2 and a low-conductance pore mutant that produced six groups ofchannel with conductance ranging from wild type to that of homomeric mutant [85].The six conductance groups were then correlated with possible arrangements ofmutant and wild-type subunits, and it was concluded that the conduction pathwayis a single pore created by an equal contribution from each subunit. A single poremodel is also consistent with the three-dimensional structure of the RyR as visualizedby single-particle electron microscopy (see below).

4. RyR structure

4.1. Three-dimensional architecture

The detailed atomic three-dimensional structure of the RyR is not known andawaits crystallization of the protein. However, gross three-dimensional reconstruc-tions have been obtained for the three mammalian RyRs at a resolution ofapproximately 30 A using image-processing techniques applied to electron micro-graphs of individual frozen-hydrated purified channels [58,86–88]. In general, theshapes of the three mammalian RyRs are very similar; there are however, somepoints of structural heterogeneity that could account for the specific properties ofeach isoform. Overall, the RyR resembles a mushroom with a ‘cap’ composed ofthe cytoplasmic domains and the ‘stalk’ made up of the membrane-spanningdomains (Fig. 1). The structure has a 4-fold cyclic symmetry around an axisperpendicular to the membrane consistent with the tetrameric nature of thechannel. The cytoplasmic part has an overall square prism shape (approximately28 nm� 28 nm� 12 nm), composed of globular parts interconnected by segments.The TM assembly has a square shape in transverse cross section, with an edgemeasuring around 12 nm at the attachment side to the cytoplasmic region and alength of about 7 nm in the direction perpendicular to the SR membrane, morethan enough to traverse a membrane bilayer. More recently, through improve-ments in the preparation of the frozen samples, software analysis and processing ofa larger number of pictures, structures have been resolved at a higher resolution(approximately 14 A) [89,90]. At the highest resolution achieved to date (9.6 A),some rod-like densities indicative of a-helices have been assigned in the mem-brane-spanning region of the channel [91].


The structure of the RyR has also been determined in an open conformation, andalthough the gross shape is similar to the closed state, there are some differences[92–94]. The channel in the open state is slightly taller because of a vertical elongationof the TM assembly and the clamp-shaped domains at the corners of the structure.Most significant is the presence of a low central density (approximately 2 nm indiameter) in the TM assembly of the open form connecting the luminal and cyto-plasmic sides, indicating a single pore within the tetrameric channel. The TMassembly is also rotated by approximately 4� with respect to the cytoplasmic region.In addition, two subdomains of the clamp region that are joined in the closedconformation of the channel are separated in the open state. These studies indicatethat alterations in the functional state of the RyR involve widespread changes in thestructure of both the cytoplasmic and TM domains of the channel.

The binding sites for two accessory proteins, calmodulin (CaM) and FKBP12,have been localized by determining differences in shape between free RyR and RyRprotein-bound complexes [95]. Both CaM and FKBP12 are situated further than10 nm away from the putative TM pore, suggesting that conformational changes canbe transmitted over long distances (Fig. 1). Several RyR regions including theN-terminal [96,97] and central domains [98] and the three divergent regions [99–102]have also been localized using chimaeras of either RyR with insertions of greenfluorescent protein (GFP) or glutathione S-transferase (GST) or binding ofsequence-specific antibodies.

4.2. Membrane topology

Molecular cloning and hydropathy plot analysis of the predicted peptide sequencehave indicated a large hydrophilic N-terminal region and a smaller hydrophobicC-terminus [16,17]. The large N-terminal region faces the cytoplasm and constitutes

Fig. 1. Ryanodine receptor three-dimensional structure. RyR architecture displayed as a projection view

from the cytoplasm to sarcoplasmic reticulum (SR) lumen (left panel), the view from SR lumen to

cytoplasm (middle panel) and side view (right panel). The numerals on the cytoplasmic assembly refer to

distinguishable globular domains. TA indicates the transmembrane assembly. The location of the three

divergent regions D1 (yellow), D2 (orange) and D3 (purple) as well as the N-terminal (red) and central

domains (light green) is shown. The location of the binding sites for FK506-binding protein 12 (FKBP12)

(brown) and calmodulin (green) is also shown. Images were kindly prepared by Dr. Zheng Liu (Wadsworth

Center, Albany, NY, USA) (See Color Plate 35, p. 533).


the ‘foot’ structure seen in electron micrographs of junctional SR. Studies of trun-cated channels have provided evidence that the C-terminal portion contains theTM sequences that form the channel pore. Limited proteolysis of the nativeRyR1 tetramer (30 S) yielded a smaller tetrameric complex (14 S) composed of theC-terminal 76-kDa fragment of the RyR1monomers, which formed a functional ionchannel upon incorporation into planar lipid bilayers [103]. Heterologous expressionof a truncated RyR1 encompassing the N-terminal 182 residues fused with theC-terminal 1030 amino acids and yielded a protein of about 130-kDa that functionedas an ion channel with conductance comparable with that of the full-length protein[104,105]. Similarly, a truncated version of an insect RyR isoform containing theN-terminal 276 residues fused with the C-terminal 1484 amino acids was also shownto function as an ion channel [106]. Cells expressing the truncated RyR1 protein hada higher resting Ca2+ concentration and smaller Ca2+ pools [104,105] that were alsoobserved in cells expressing RyR2 C-terminal constructs [107]. These findings suggestthat the RyR C-terminus forms a Ca2+ conduction pore that is constitutively open.

Putative membrane-spanning domains are located in the C-terminal (10–20% ofthe protein, last 500–1000 amino acids), and two topology models of 4TM or 10TMsegments have been proposed [16,17]. Based on their hydrophobicity and theirconservation across species and isoforms, the four segments of the 4TM modelcorresponding to M5, M6, M8 and M10 of the 10TM model emerge as strongcandidates for membrane-spanning helices. Both models are consistent with thecytoplasmic location of the amino- and carboxyl-termini shown by immunolocaliza-tion studies, indicating that an even number of sequences traverse the membrane[108,109]. The further use of site-directed antibodies discriminating between cyto-plasmic and luminal locations was in support of the 4TM model although incon-sistent with the assignment of M9 as a membrane-spanning sequence with thecorresponding M8–M10 loop found to be luminal [39,109].

The use of C-terminal-truncated channels carrying a C-terminal GFP tag, withdeletions starting near the beginning or the end of predicted TM helices M1–M10,indicated that RyR1 contains eight TM sequences organized as four hairpin loopsformed from M4a–M4b, M5–M6, M7a–M7b and M8–M10 [110,111]. Hydrophobicsegments M1, M2, M3 and M9 did not traverse the membrane, whereas the long(approximately 50 residues) M7 domain was shown to form a double TM hairpin.M4 was found to be membrane bound, but the mechanism of membrane associationis unclear; M4 can either form a TM hairpin or associate in an unorthodox fashionwith the cytosolic leaflet of the membrane. Support for the above biochemicalfindings is provided by the most recent three-dimensional reconstruction of theRyR1 at 9.6 A resolution using single-particle electron microscopy of frozen purifiedchannels [91]. Five helices were clearly resolved in the membrane-spanning portion ofthe channel, although additional helices may exist that could not be unambiguouslyidentified at the resolution achieved. However, the structure obtained suggested amore complicated arrangement than a simple arrangement of membrane-spanninghelices, because only helix 1 (assigned to the predicted M10 domain) had eithersufficient length (approximately 45 A) or the proper orientation to span the mem-brane. Helix 3 was found oriented parallel to the plane of the membrane on the


cytoplasmic side and most probably corresponds to M4. The currently favoured RyRmembrane topology is depicted in Fig. 2.

4.3. Pore structure

The determination of the structure of K+ channels at atomic resolution [112,113] hasenabled the prediction of critical sequences in RyR that participate in cation trans-location and selection. The conserved GGGIG motif in RyRs (residues 4894–4898 inRyR1 within domain M9) is a sequence related to the K+ channel TVGYG motifthat comprises the selectivity filter implying that the luminal loop between the lasttwo TM helices (M8 and M10) may fold back into the membrane in a manner similarto that found for the K+ channel [114]. Several studies of recombinant mutant RyR1or RyR2 channels using measurements of intracellular Ca2+ concentration, ryano-dine binding and single channel recordings have indicated that mutations within theGGGIG motif and neighbouring residues produce profound alterations in channelactivity and, most importantly, in cation conductance and selectivity [85,115–117].Channel activation and gating were also altered by mutations in the last TM helix

Foot region

Cytosol

4911

4854

48004803

4637

4576

4935

48344825

4776

4662

455743

714322

Pore loop

5037

Lumen

M4

M5 M6 M7 M8 M10

Fig. 2. Ryanodine receptor (RyR) membrane topology. Transmembrane domain assignment is based on

the 10 transmembrane (10TM) model [17] and revised according to recent experimental evidence

[91,110,111]. Hydrophobic sequences M5, M6, M8 and M10 are shown to span the membrane once, M7

is shown as a transmembrane hairpin and M4 to reside parallel to the membrane on the cytoplasmic side.

Hydrophobic domain M9 dips inside the lipid bilayer from the luminal side and forms the pore loop and

selectivity filter. The bulky amino-terminal portion forms the ‘foot structure’ seen in electron micrographs.

Coordinates given are for rabbit RyR1.


suggesting that it forms the inner helix of the pore [118–120]. In addition, otherstudies have identified the presence of rings of negative charges in the luminal (D4899and E4900) and cytosolic (D4938 and to a lesser extent D4945) vestibules that arerequired for maintaining high rates of ion translocation and Ca2+ selectivity in RyR1[121,122]. It has also been proposed that the conserved GLIIDA motif within M10(residues 4934–4939 in RyR1) is analogous to the gating hinge motif (GXXXXA)identified in K+ channels, with G4934 (G4866 for RyR2) introducing a kink in theTM helix [82,91,122]. The importance of domains M9 andM10 in channel function isalso highlighted by the occurrence of natural mutations associated with skeletal andcardiac muscle diseases (see Sections 10.2. and 10.3.).

The pore structure of the closed RyR1 channel has been described at 9.6 Aresolution, the highest resolution achieved to date [91]. At this resolution, at leastfive helices within the membrane-spanning region could be resolved, including TMhelix 1 that was noticeably kinked and short helix 2 both lining the pore. Helix 1,which is likely to correspond to M10, was assigned as the inner, pore-lining helix,whereas helix 2, likely to correspond to M9, was assigned as the pore helix associatedwith the selectivity filter. The arrangement of the helices and the kink in the inner,pore-lining helix suggested that the RyR structure resembles the pore structure of theMethanobacterium thermoautrophicum K+ channel. A hypothetical model of theRyR ion pore is shown in Fig. 3.

5. E–C coupling

E–C coupling is a term used to describe the events that link plasma membranedepolarization to the release of Ca2+ from the SR, which in turn triggers contraction.Central to this process is the functional interaction between the RyR and the surfacevoltage-activated L-type Ca2+ channel or dihydropyridine receptor (DHPR).

In skeletal muscle, the terminal cisternae (TC) of the SR (also referred to asjunctional SR) are closely apposed to either the surface membrane to form the‘peripheral couplings’ or the transverse tubules (T-tubules), which are tubular inva-ginations of the plasma membrane, to yield the ‘triads’ (reviewed in [2]). The triad isso named because it is composed of two TC closely facing a central T-tubule. RyR1sare visible in electron micrographs as dense particles initially named ‘feet’ that spanthe whole distance between the junctional SR and the surface membrane. Electronmicroscopy studies have also indicated that RyR1s are concentrated on the TC andform highly ordered arrays [123,124]. The skeletal muscle DHPR isoform (DHPRs)on the plasma membrane and T-tubules in particular acts as voltage sensor as well ascalcium channel, and it is coupled through a direct physical association with RyR1[125–130]. This direct mechanical coupling provides the basis of the E–C couplingmechanism in skeletal muscle (Fig. 4). Thus, T-tubule depolarization is sensed by theDHPRs that assumes a conformational change that is then transmitted to RyR1causing the channel to open and release Ca2+ from the SR. Ca2+ influx is very small,and in fact, adult vertebrate skeletal muscles can contract vigorously even in theabsence of extracellular Ca2+.


Morphological studies have also shown that DHPRs channels are clustered ingroups of four (‘tetrads’), which form ordered arrays that face the ordered RyR1arrays on the SR [131–133]. The formation of DHPRs tetrads is dictated by theirinteraction with RyR1 in such a way that each of the four DHPRss composing thetetrad is linked to a subunit of an underlying RyR1. Interestingly, DHPRs tetrads areassociated with alternate RyR1s, leaving a set of orphan RyR1s that are not directlylinked to DHPRss. It is believed that uncoupled RyR1s are either activated by Ca2+

released from adjacent, DHPRs-coupled channels or by transmission of the confor-mational change through physical interactions between RyR1s within the array. Thefunctional interaction between RyR1 and DHPRs is bidirectional because DHPRs

controls gating of RyR1, and the RyR1 in turn affects DHPRs channel properties[134].

In cardiac muscle, the SR has a similar but not identical arrangement to that ofskeletal muscle (reviewed in [2]). Peripheral couplings are also found in cardiomyo-cytes, but the structures corresponding to the triads are ‘diads’, because TC arecomparatively less numerous, occupying only one of the two faces along the

Cytosol

Asp 4945

Asp 4899Glu 4900

Selectivityfilter

Asp 4938

Gly 4934

Pore-lining helix

G G

GG

G G

GG

I I

Fig. 3. Ryanodine receptor (RyR) pore structure. Hypothetical model of the RyR pore structure based on

the model proposed by [122]. The inner transmembrane helix corresponds to the predicted M10 domain,

whereas the pore helix corresponds to M9 with the GGGIG motif (residues 4894–4898 in RyR1) forming

the selectivity filter. The putative position of negatively charged amino acids D4899 and E4900 on the

luminal side and D4938 and D4945 on the cytosolic side is shown. G4934may act as a gating hinge

producing a kink in the inner transmembrane helix. Note that only two of the four pore-forming

segments are shown. Image kindly prepared by Dr. Leon D’Cruz (Wales Heart Research Institute, Cardiff

University, UK).


T-tubules. Moreover, TC, termed corbular or extended SR, are also present in theinterior of the cell without bearing any relationship to the plasma membrane. Incardiac muscle, although the cardiac DHPR isoform (DHPRc) and RyR2 are loca-lized in close proximity to each other, there is no evidence for direct physicalinteraction, and the DHPRc does not form ordered arrays of tetrads but they arerandomly distributed with respect to the RyR2 arrays [135–137]. There is however, afunctional DHPRc–RyR2 association, and Ca2+ influx is an absolute requirementfor the mechanism of E–C coupling in the heart [128,138–141]. Thus, plasma mem-brane depolarization activates the DHPRc allowing extracellular Ca2+ to flow intothe cytoplasm, which in turn activates RyR2 channels to release Ca2+ from the SR(Fig. 4). It has been estimated that the opening of a single DHPRc channel triggersCa2+ release from 4–6 RyR2 channels [118]. This mechanism is known as ‘calcium-induced calcium release’ (CICR), and in the case of the heart, this term refers toRyR2 activation by Ca2+ coming from both outside the cell and within the SR. Thespatial organization of DHPRc with relation to RyR2 channels is very important fornormal E–C coupling. Reduced efficiency of E–C coupling has been implicated in

Skeletal musclemechanical coupling

Plasmamembrane

Voltage-gated Ca2+ channel

(DHPRC)

RyR1 RyR2

Sarcoplasmic reticulum Sarcoplasmic reticulum

Ca2+ Ca2+ Ca2+

Ca2+

Ca2+

Plasmamembrane

Voltage sensor(DHPRS)

Cardiac musclecalcium-induced calcium release

Fig. 4. Ryanodine receptor (RyR) activation in striated muscles. In skeletal muscle dihydropyridine receptor

(DHPRs) channels sense the plasma membrane depolarization and transmit a conformational change to the

RyR1 causing the channel to open and release Ca2+ from the sarcoplasmic reticulum (SR) store. Ca2+ influx

is not required. Alternate RyR1 channels that are not coupled to DHPRs may either be activated by Ca2+

released from a neighbouring RyR1 or by transmission of the conformational change through physical

interactions between RyR1 channels within the array. In cardiac muscle, DHPRc channels are activated by

plasma membrane depolarization and allow Ca2+ influx into the cytoplasm. The Ca2+ entering the cell

activates RyR2 channels allowing Ca2+ release from the SR store. Neighbouring RyR2 channels may in turn

be activated by Ca2+ flowing fromwithin the SR or by conformational changes transmitted through physical

interactions between RyR2 channels within the array (See Color Plate 36, p. 533).


heart failure (see Section 10.1.), attributed to a reduction in the density of T-tubulesand therefore DHPRc channels and/or reorganization of T-tubules resulting in agreater number of functionally uncoupled RyR2 channels [142–146].

The CICR process is intrinsically self-regenerating because the Ca2+ released by aRyR2 channel should feedback and further activate the channel or its neighbours, andit should therefore lead to release of all the calcium stored in the SR. However, SRCa2+ release in an all-or-none fashion is not observed, but rather it is graded andtightly controlled by the magnitude and duration of the L-type Ca2+ current. Theimplication is that some sort of negative control mechanism(s) must exist to counterthe inherent positive feedback of CICR and terminate calcium release. Several mechan-isms have been proposed, including Ca2+-dependent RyR inactivation that may alsoinvolve accessory protein(s), stochastic attrition, adaptation, coupled gating of adja-cent channels and local depletion of SR calcium; however, none of these mechanismsappears sufficient by itself to terminate calcium release (reviewed in [147,148]).

6. Modulation by pharmacological agents

A plethora of exogenous compounds, including volatile and local anaesthetics,4-chloro-m-cresol, polylysine, doxorubicin, dantrolene, neomycin and peptide toxins,has been shown to modulate RyR channel activity (reviewed in [149]). Of special noteis caffeine, which is the most commonly used drug to activate the RyR, rutheniumred, which is the most commonly used channel blocker, and ryanodine, which isdiscussed Section 6.1.

6.1. Ryanodine

Ryanodine and related ryanoid compounds have complex effects on channel con-ductance and gating in all three RyR isoforms (reviewed in [3]). Initial studies usingskeletal and cardiac muscle preparations demonstrated that ryanodine results in anincrease as well as in a decrease in the Ca2+ permeability of the SR, and its effects areconcentration and use dependent. In particular, ryanodine at low concentrationsactivates Ca2+ release from the SR but prolonged exposure to micromolar concen-trations inhibits Ca2+ release.

The action of ryanodine is clearly documented by single channel recordings[66,150–152]. Several studies have shown that ryanodine at submicromolar to micro-molar concentrations (up to 10mM) results in the channel exhibiting partially conduct-ing states. Multiple subconductance levels have been reported but the most frequentlyobserved one is a sole, long lasting substate that is approximately 40–50% of the fullconductance state. This effect is observed consistently and has become a signature for aryanodine-modified RyR channel. Ryanodine at very low doses (approximately 10nM)increases the frequency of channel openings to the full conductance level, whereasmicromolar concentrations (>10mM) block the channel. Further studies demonstratedthat ryanodine sensitizes the RyR to Ca2+ activation through an allosteric mechanism


of interaction [153–155]. It was also shown that the ryanodine-modified channel issensitive to Mg2+ inhibition and to activation by caffeine and ATP.

The complex effects of ryanodine are most likely due to the presence of both high-affinity (Kd� 10 nM) and low-affinity (Kd> 1mM) sites [8,156–158]. Resultsobtained for [3H]–ryanodine binding to purified RyRs suggest a single high-affinitysite per tetrameric RyR channel. Occupation of the low-affinity sites slows thedissociation of ryanodine from the high-affinity site. After occupation of the low-affinity sites, the channel undergoes a slow transition to a state characterized bypersistent channel inactivation, which is associated with decreased ryanodine bind-ing. It has been proposed that these effects are due to an allosteric or steric negativeinteraction between four initially identical binding sites, with high-affinity bindingfor the first ryanodine molecule locking the channel in a subconductance state andinhibiting ryanodine binding to the other sites. High-affinity ryanodine binding isobserved under conditions that are associated with activation of the RyR channel,indicating that ryanodine binds to a conformation associated with the open state ofthe channel [156,159]. This property has proved useful, because it allows ryanodinebinding to be used as an index of RyR channel activation.

The high-affinity binding site for ryanodine has been localized to the carboxyl-terminal domain of the protein. Limited proteolysis of the native RyR1 tetramer (30 S)that was covalently bound with [3H]–ryanodine (with the use of a photo-activatedcross-linking derivative) or pre-labelled with it yielded a smaller tetrameric complex(14 S) composed of the C-terminal 76-kDa fragment of the RyR1monomers, whichretained pre-bound [3H]–ryanodine [160,161]. Further evidence comes from expressionstudies of truncated, C-terminal RyR protein fragments that formed functional chan-nels upon incorporation into planar lipid bilayers and were modified to the character-istic approximately 50% subconductance state by ryanodine [104,106]. It has beensuggested that ryanodine binding occurs within the channel pore, as mutation ofresidues within the pore region (M9 andM10 domains) alters the ability of the receptorto bind ryanodine without affecting other characteristics of channel function [117,118].Moreover, interaction of ryanodine and related ryanoids with the RyR is voltagedependent and sensitive to the net charge of the ryanoid, suggesting that the site ofinteraction may be within the voltage drop across the channel pore [162,163].

7. Modulation by endogenous effectors

RyR channel activity is regulated by a wide variety of endogenous molecules, withCa2+, Mg2+ and ATP being the key regulators (reviewed in [149]).

7.1. Cytosolic Ca2+

Ca2+ is thought to be the physiological channel activator because other ligands eithercannot activate the channel in its absence or they require Ca2+ for maximum effect.SR Ca2+ flux measurements, single channel recordings and ryanodine-binding assays


have demonstrated that cytosolic Ca2+ has a biphasic effect on skeletal muscle RyRchannel activity [164–169]. The threshold for channel activation is approximately100nM with a maximum in the range of 10–100mM, whereas millimolar Ca2+ almostentirely inhibits the channel. Such a bell-shaped, cytosolic Ca2+-dependence curvesuggests that the RyR1 contains high-affinity Ca2+-binding site(s) that stimulate thechannel and inhibitory low-affinity site(s). However, Ca2+ alone cannot fully activateRyR1. The cardiac RyR shows some marked differences to the skeletal type [170–175].The most important ones are that RyR2 is less sensitive to inhibition by high Ca2+

concentrations and that almost maximum activation can be achieved by Ca2+ alone atapproximately 100mM. RyR3 resembles RyR2more closely than RyR1, with lowersensitivity to Ca2+ inactivation and full activation by Ca2+ alone at approximately100mM [58,70,167,168,176]. It has also been reported that RyR1 and to a lesser extentRyR2 channels are functionally heterogenous with respect to Ca2+ regulation[177,178], which may be due to differences in the redox state between individualchannels [179]. Characterization of the three mammalian, recombinant RyRsexpressed in a non-myogenic cell line (HEK293) revealed cytosolic Ca2+-dependenceproperties similar to the native channels [22,25,180].

7.2. Luminal Ca2+

The rate of Ca2+ release from the SR is sensitive to the store Ca2+ content [181,182],and single channel recordings have demonstrated effective RyR regulation by luminalCa2+ [183–188]. Although differences may exist between RyR1 and RyR2, increasingluminal Ca2+ concentration augments channel activity by increasing the channelsensitivity to agonists such as cytosolic Ca2+, ATP and caffeine and by alleviatingMg2+ inhibition. However, increasing luminal Ca2+ levels above a threshold causes adecrease in channel activity. The mechanism of RyR activation by luminal Ca2+ islikely to involve either Ca2+ regulatory site(s) on the luminal side of the RyR[183,184,187] and/or the cytosolic Ca2+ regulatory site(s) following permeationthrough the pore [185,186]. It has been proposed that both mechanisms are opera-tional, with luminal Ca2+ altering the affinity of cytosolic regulatory site(s) through anallosteric effect in single isolated channels, whereas in close-packed RyR1 arrays,luminal Ca2+ flowing through one channel interacts directly with cytosolic regulatorysite(s) of neighbouring channels [188]. The luminal Ca2+ sensor can either be anintrinsic property of the RyR or is imparted by accessory protein(s). Evidence forthe former is provided by proteolysis studies suggesting the presence of both activatingand inactivating Ca2+ sites on the RyR luminal side [189]. In contrast, dissociation ofcalsequestrin (CSQ), triadin and junctin abolished RyR2 regulation by luminal Ca2+

[190], whereas CSQ association amplified channel responses to luminal Ca2+ [191].

7.3. Mg2+

Mg2+ is a potent RyR channel inhibitor. SR Ca2+ flux measurements, single channelrecordings and ryanodine-binding assays have demonstrated that Mg2+ inhibits


RyR1 in a dose-dependent manner, with millimolar concentrations resulting incomplete inactivation [164–166,188,192–194]. Mg2+ inhibition may be a significantmechanism for maintaining the RyR1 closed as the cytoplasmic concentration isapproximately 1mM. RyR2 and RyR3 are also sensitive to Mg2+ inhibition but to alesser extent [25,70,170,194–196]. It has been suggested that Mg2+ exerts its inhibi-tory effects by competing with Ca2+ for the high-affinity Ca2+-stimulatory site(s)and by binding to the low-affinity inactivation site(s) with an affinity comparablewith Ca2+ [193,194,196]. More recent evidence suggests that Mg2+ is an antagonistthat inhibits RyR1 channel activity in the absence of Ca2+ [188].

7.4. ATP

The adenine nucleotides ATP, ADP, AMP and cyclic-AMP, as well as adenine,activate RyR channel activity [164–166]. SR Ca2+ flux measurements, single channelrecordings and ryanodine-binding assays have demonstrated that ATP at millimolarlevels is a potent agonist of skeletal muscle RyR [164–166,197,198]. RyR1 is stimu-lated by ATP even at very low nanomolar Ca2+ concentrations, whereas the combi-nation of micromolar Ca2+ and millimolar ATP elicits persistent channel activation.It has been suggested that ATP increases the sensitivity of the channel to Ca2+

activation and decreases its sensitivity to Ca2+ inactivation [169]. The effect ofATP on the cardiac RyR is qualitatively similar although less marked[170,195,199]. ATP also stimulates RyR3 channel activity and overrides inhibitionby millimolar Ca2+ [25,200].

7.5. Redox status

Pharmacological sulphydryl-reactive reagents, including thimerosal, dithiodipyri-dines, N-ethylmaleimide and diamide, have been shown to activate skeletal andcardiac muscle RyRs [201–203]. This effect was reversed by reducing reagents suchas dithiothreitol, whereas prolonged exposure to oxidants, or use of high concentra-tions, led to irreversible loss of channel activity. Endogenous physiological redoxmodulators have also been shown to regulate the RyR. Glutathione in its reducedform was found to inhibit the channel (but oxidized glutathione was stimulatory)[204,205], whereas reactive oxygen species such as hydrogen peroxide and superoxideanion radical activated the channel [206–209]. Reactive oxygen species and super-oxide anion radical in particular, generated by an endogenous SR NAD(P)H oxi-dase, have been implicated in modulation of skeletal and cardiac RyRs [209–212].Nitric oxide has also been proposed to be an important physiological modulator ofRyR function, although results obtained from different studies are contradictory[213–218]. Native RyRs are endogenously S-glutathionylated as well as S-nitrosy-lated [212,215,216,219], suggesting that these post-translational modifications mayplay a role in RyR function.

The redox potential of RyR1, estimated to be about –165mV, was found to besensitive to channel modulators [205]. Conditions that favour channel opening


lowered RyR1 redox potential that favours the oxidation of critical thiols, whereasconditions that close down the channel have the opposite effect. There is evidence forat least three classes of functional cysteines that modulate RyRs, and channel activityhas been correlated with the number of free thiols [202,215,220]. Under physiologicalconditions, RyR1-free thiol content is high (approximately 48 per subunit) and isassociated with low channel activity. Oxidation of approximately 10 thiols has littleeffect on channel activity, whereas oxidation of approximately 15 extra thiolsincreased channel activity and was reversible. More extensive oxidation (loss ofapproximately 10 additional thiols) inactivated the channel irreversibly.

RyR sulphydryl modification alters Ca2+ dependence and increases the sensitivityof the channel to Ca2+ activation [179,205,220–222]. In addition, it was shown thatRyR oxidation increases the sensitivity of the channel to ATP activation [221,222]and decreases its sensitivity to Mg2+ inhibition [217,223]. Sulphydryl modification ofRyR involves altered protein–protein interactions, in particular formation of intra-subunit and intersubunit cross-links within RyR1 [224], high-molecular weight com-plexes with triadin [225], CaM binding [208,216,219,226] and FKBP12 binding [219].Notably, defective regulation of RyR2 channels due to oxidative stress has beenimplicated in heart failure [227] (see Section 10.1.).

7.6. Phosphorylation status

In vitro phosphorylation studies have shown that both skeletal and cardiac RyRs aresubstrates for serine/threonine protein kinases, primarily for cAMP-dependent pro-tein kinase (PKA) and Ca2+/calmodulin-dependent protein kinase II (CaMKII) andless efficiently for cGMP-dependent protein kinase (PKG) [228–233]. Dephosphor-ylation is carried out primarily by protein phosphatases 1 (PP1) and 2A (PP2A)[234,235]. CaMKII is tightly bound to SR membranes and co-purifies with the RyRthrough a direct association [229,232,236]. RyR association with PKA, PP1 andPP2A appears to be indirect, through specific anchoring proteins [235]; this explainswhy endogenous CaMKII but not PKA (or PKG) activity is often reported in isolatedSR membranes [229,230,232,237–239]. Although results vary, much evidence suggeststhat there are multiple RyR sites phosphorylated by CaMKII [230–233,240–242].Two in vivo phosphorylation sites have been identified: RyR2 S28081 and thecorresponding RyR1 S2843, whereas a second site, S28141, has been found for RyR2only.

Studies aiming to identify the PKA site(s) have produced conflicting results overthe issue of one or two sites and which one is PKA specific. Although RyR2 S2808was initially reported to be CaMKII specific [231], it was later proposed that thisresidue (and the corresponding RyR1 S2843) is phosphorylated exclusively by PKA[234,241,243]. However, recent studies have identified a second PKA site, S20311 inaddition to S2808, and both appeared to be phosphorylated in vivo [242,244].Expression of mutant channels, where these serines were substituted by alanines(which cannot be phosphorylated) either individually or in combination, did notresolve the discrepancy. One group observed that single S2808A substitution (but not


S2031A) abolished PKA-dependent phosphorylation [245,246], whereas anothergroup demonstrated PKA-dependent phosphorylation occurred with either indivi-dual mutant RyR2 and was absent only for the double mutant [242,244]. Determina-tion of S2808 phosphorylation characteristics is important because it has beenimplicated in the pathogenesis of heart failure (see Section 10.1.). Many studieshave indicated that S2808 is phosphorylated to high levels (up to 75%) even innormal hearts at rest and can be phosphorylated efficiently by several protein kinasesincluding PKA, CaMKII and PKG [231,233,240,242,244,247,248].

The functional effects of RyR phosphorylation/dephosphorylation have beenexamined by SR Ca2+ flux measurements, single channel recordings and byryanodine-binding assays. Addition of exogenous PKA results in increased RyRactivity because of enhanced sensitivity to Ca2+ activation and substantially reducedMg2+ inhibition [228,230,234,237,238,243,248–252]. These effects were reversed byPP. One study reported that PKA activated the RyR but only during rapid rises inlocal Ca2+ concentration, whereas under fixed Ca2+ levels, it promoted channelclosure [249]. Another study has suggested that the PKA-stimulatory effects occuronly when stoichiometric phosphorylation of S2808 is achieved; phosphorylation byup to 75%, as often is the case in normal hearts at rest, had negligible effects onchannel activity [248]. Exogenous CaMKII displayed enhanced RyR channel activity[231,236–238,241,250] although one study reported inhibition [253]. Contradictoryresults were also obtained when the effects of endogenous CaMKII (activated byCa2+ and/or CaM) were assessed [230,237–239]. The reasons for these discrepanciesare unknown. Notably, PKA- and CaMKII-dependent phosphorylation produceddifferent functional effects in parallel investigations, suggesting that they targetdifferent residues and/or a different number of residues [230,237,238,250]. Depho-sphorylation of RyRs is expected to inhibit channel activity because phosphorylationresults in channel activation (in the majority of studies). However, RyRs treated withPP displayed increased channel activity [248,253,254]. These apparently disparatefindings are explained when the basal RyR phosphorylation level is considered. It hasbeen suggested that phosphorylation levels above (i.e. additional phosphorylation)or below (i.e. dephosphorylation) an intrinsic homeostatic level results in channelactivation [248].

In cardiomyocytes, b-adrenergic stimulation leads to cAMP elevation and PKAactivation, resulting in enhancement of E–C coupling (reviewed in [255]). This isdue to increased DHPR Ca2+ current, as well as increased SR Ca2+ content causedby higher SR Ca2+-ATPase pump activity following relief of phospholambaninhibition. RyR activity may also be up-regulated following b-adrenergic stimula-tion as in vitro functional assays suggest. In support of this hypothesis, one studyfound enhancement of highly localized Ca2+ release events despite lower SR Ca2+

content [256]. However, other studies did not concur with an effect throughPKA-mediated phosphorylation. b-Adrenergic stimulation was found to have noeffect on Ca2+ transients and E–C coupling (when normalized for DHPR Ca2+

current and SR Ca2+ content), although it did enhance the initial rate of SR Ca2+

release [257]. It was further shown that direct PKA activation, upon addition ofcAMP or PKA catalytic subunit in permeabilized cardiomyocytes of


phospholamban knock-out mice, did not affect Ca2+ spark frequency (underconditions that excluded PKA effects on DHPR Ca2+ current and SR Ca2+

content) [258,259]. Interestingly, several studies have indicated that CaMKIIenhances E–C coupling by directly acting on the RyR. Activation of endogenousCaMKII in cardiomyocytes augmented electrically evoked Ca2+ transients (whennormalized for DHPR Ca2+ current and SR Ca2+ content) and increased Ca2+

spark frequency and duration (under conditions that excluded CaMKII effects onDHPR Ca2+ current and SR Ca2+ content), whereas CaMKII inhibition decreasedCa2+ spark frequency [236,259,260]. CaMKII (dC isoform) over-expression,chronically in transgenic mice or acutely in isolated cardiomyocytes through ade-novirus, enhanced electrically evoked Ca2+ transients and Ca2+ spark frequency(normalized for SR Ca2+ content) [261,262]. One study reported disparate results,where constitutively active CaMKII reduced and CaMKII inhibition enhancedCa2+ release [263]. Notably, PKA- and/or CaMKII-dependent phosphorylationof RyR2 has been implicated in heart failure (see Section 10.1.).

The effects of PP1 and PP2A in cardiomyocytes are unclear at present. Intracel-lular dialysis of PP1 and PP2A was found to decrease Ca2+ transients and E–Ccoupling without affecting SR Ca2+ content [264]. In contrast, in permeabilizedcardiomyocytes PP1 and PP2A initially enhanced Ca2+ spark frequency followedby a subsequent decline because of depletion of the SR calcium store [254].

8. Functional interactions within the RyR

8.1. Physical coupling between channels

A characteristic feature of RyRs is their membrane organization into ordered arrays,observed in both skeletal and cardiac muscle [123,124,137]. Such a regular organiza-tion is an intrinsic property of the RyR, because recombinant RyR1 expressed in anon-myogenic (CHO) cell line formed extensive arrays [265] and purified RyR1 wasshown to self-assemble into large two-dimensional crystalline arrays in a strict ionicstrength-dependent manner [266,267]. In these ‘checkerboard-like’ lattices, eachRyR1 is interlocked with four adjacent channels through a specific intermoleculardomain–domain interaction that appears to involve sequences in close proximity towhere the D2 region has been localized [102,267]. Biochemical evidence has impli-cated a sequence (residues 2540–3207) in the central region of RyR2 participating intetramer–tetramer interactions [268]. Physical coupling between RyRs may facilitateintermolecular allosteric interactions and conformational changes transmitted fromone RyR to four adjacent channels. This inter-RyR physical coupling mechanismmay explain how skeletal muscle RyR channels that are not associated with DHPRvoltage sensors can be regulated or how a given number of RyRs within a clusteropen and close simultaneously during a Ca2+ spark. Ca2+ sparks are small localizedCa2+ release events generated by spontaneous openings of RyR channels (reviewedin [269]). Ca2+ sparks are widely used as an index of intrinsic RyR activity in cell-based assays.


8.2. Interdomain interactions within the tetrameric channel

The RyR carboxyl-terminus contains the membrane-spanning domains, is capable ofself-association and functions as a Ca2+-conducting pore (see Section 4.2.). Thecytoplasmic C-terminal tail is important in channel oligomerization as a truncatedRyR1 protein lacking the last 15 residues formed inactive channels due to impairedassembly of the tetrameric complex [270], whereas the RyR2 extreme C-terminal 100residues form dimers and tetramers [271]. The large N-terminal cytoplasmic portionremains associated with the TM assembly in partial protease digestion studies ofRyR1 [272]. This is consistent with complementation assays demonstrating thatN-terminal RyR2 fragments can combine with overlapping C-terminal fragmentsencompassing the pore-forming TM domains to form functional channels [107,273].

An important role for the central domain in channel regulation and a putativeinteraction with the N-terminal region has been proposed [274–279]. SR Ca2+ fluxmeasurements, single channel recordings and ryanodine-binding assays showed thata synthetic peptide termed DP4 (residues 2442–2477 of RyR1), as well as anti-DP4antibody, enhances RyR1 channel activity and potentiates agonist-induced Ca2+

release. DP4 also enhanced Ca2+ release and contraction in skeletal muscle fibres,overriding Mg2+ inhibition, and increased the frequency of Ca2+ sparks. DP4 wasshown to bind the approximately 150-kDa N-terminal RyR1 calpain fragmentsuggesting that the entral domain interacts with the N-terminal region. Interestingly,when DP4 had a single residue change to mimic a malignant hyperthermia (MH)mutation (R2458C) (see Section 10.3.), the resulting peptide lacked any stimulatoryeffects. Similarly, an RyR2 central domain peptide (DPc10, residues 2460–2495) wasfound to activate RyR2, an effect that was abolished by a single residue change(R2475S) mimicking a mutation involved in catecholaminergic polymorphic ventri-cular tachycardia (CPVT) (see Section 10.2.) [280]. Furthermore, DPc10 decreasedCa2+ transients and cell shortening due to Ca2+ leak from the SR calcium storewhile transiently increasing Ca2+ spark frequency in cardiomyocytes [281,282]. LikeDP4, DPc10 was also shown to bind the approximately 120-kDa N-terminal calpainfragment of RyR2. Hence, stimulatory effects of central DPcs may be due to disrup-tion of the normal interdomain interactions that stabilize the closed state of thechannel, with DP4 or DPc10 competing with the corresponding domain of the nativeRyR for a binding site on the N-terminal region. However, other studies appear to beinconsistent with this hypothesis. A RyR1N-terminal fragment (residues 281–620)interacted with two native RyR1 fragments (residues 799–1172 and 2937–3225) thatdo not encompass the DP4 sequence [283]. A RyR2N-terminal peptide termed DP1-2 (residues 601–639) and the corresponding RyR1 peptide (residues 590–628) bothstimulated channel activity of the cardiac RyR but had no effect on skeletal RyRfunction [284]. Further peptide-probe studies demonstrated that dantrolene, whichinhibits Ca2+ release from skeletal but not from cardiac muscle, binds native RyR1at residues 590–609 (DP1 peptide) and inhibits the stimulatory effect of DP4. How-ever, dantrolene does not bind native RyR2 although it does bind the identicalcardiac DP1 peptide (residues 601–620) [285–287]. Similarly, anti-DP1 antibodyrecognizes native RyR1 but not native RyR2. In additon, DP4 was reported to


have no effect on the type 3 RyR [288]. These latter studies suggest that thepostulated N-terminal interaction with the central domain may be isoform specific.

The primary CaM-binding site (see Section 9.3.) has also been implicated in inter-domain interactions [289–292]. A synthetic peptide comprising the CaM site (aminoacids 3614–3643) activated RyR1 channel activity in SR Ca2+ flux measurements,single channel recordings and ryanodine-binding assays and increased Ca2+ sparkfrequency in skeletal muscle fibres, effects which were CaM independent. Further-more, RyR1mutant channels lacking this region displayed a severely compromisedagonist-induced Ca2+ release. Peptide 3614–3643 were able to bind to a region(amino acids 4064–4210) with structural similarities to CaM in a Ca2+- and CaM-dependent way. Thus, it was proposed that an interdomain, CaM-sensitive interac-tion occurs between the CaM-binding site and the CaM-like domain, which stabilizesthe channel in a closed conformation.

Evidence also exists for intersubunit contacts between cytoplasmic domains ofRyR1, from a combination of protease-protection and cysteine-modification assays[202,224,293,294]. These studies demonstrated that an intersubunit disulphide bondcould be formed by treating RyR1 with oxidizing reagents, involving C3635 withinthe primary CaM-binding site and a second cysteine located within residues 2000–2401.An additional intrasubunit disulphide bond was also detected, but the cysteinesinvolved remain to be identified.

9. RyR accessory proteins

The RyR is a dynamic macromolecular complex interacting directly or indirectlywith numerous proteins that affect its channel function, including protein kinasesand phosphatases docked through anchoring proteins [234,235], S100 [295], calexci-tin [296], homer [297] and snapin [298]. The DHPR, FKBP, CaM, sorcin, CSQ,triadin and junctin are the best characterized RyR accessory proteins and arediscussed Section 9.1.–9.5.

9.1. DHPR

The DHPR is the L-type plasma membrane Ca2+ channel that senses surfacemembrane depolarization and initiates the process of E–C coupling in striated muscle(see Section 5.). The DHPR is composed of a1-, a2-d-, b- and g-subunits, with a1

constituting the channel-forming, voltage-sensing and dihydropyridine-sensitive por-tion of the receptor. The skeletal muscle a1s-subunit is composed of 1873 aminoacids, with a predicted topology consisting of four repeats of six TM segments linkedby cytoplasmic loops of variable length bounded by cytoplasmic N- and C-termini[299]. The cardiac muscle a1c-subunit displays 66% homology with the skeletalisoform, with similar TM topology but longer N- and C-terminal domains [300].

Expression of DHPR isoforms in skeletal myocytes derived from the dysgenicmouse, a naturally occurring mutant that lacks functional DHPR a1-subunit,


demonstrated that a1s restores depolarization-induced intracellular Ca2+ transientsin the absence of Ca2+ influx, i.e. skeletal type E–C coupling, whereas expression ofa1c results in cardiac type E–C coupling (dependent on Ca2+ influx) [127,139].Analysis of skeletal and cardiac isoform chimaeras of the a1-subunit indicated thatthe major determinant of skeletal E–C coupling is the II–III cytoplasmic loop of thea1s-subunit [128]. Further studies of additional chimaeras, or mutant a1s-subunitswith scrambled sequences, deletions or point substitutions, identified residues720–764 within the II–III loop (also known as peptide C) to be crucial for skeletal-type E–C coupling [301–305]. The minimal region required was found within residues725–742, with four individual residues being particularly important (A739, F741,P742 and D744).

The recombinant a1s-subunit II–III loop activated RyR1 channels in ryanodine-binding assays and single channel recordings [306,307]. Further studies to identify theminimal region within the II–III loop affecting RyR1 channel activity have producedconflicting results. Synthetic peptide C (or a larger fragment containing this region)was reported to have no effect, inhibit or activate the RyR1, depending on experi-mental conditions (e.g. peptide concentration, incubation time and effectors such asCa2+, Mg2+ or ATP) [279,308–315]. A second peptide (termed peptide A) corre-sponding to residues 671–690 (or a larger fragment containing this region) activatedRyR1 in SR Ca2+ flux measurements, single channel recordings and ryanodine-binding assays [308,309,311,312,316–319]. These studies showed that a basicsequence RKRRK is essential for the action of peptide A and that phosphorylationof S687 could also be involved in RyR1 activation. Peptide A activation of RyR1 wasalso dependent on the presence of FKBP12. Interestingly, peptide C suppressed thestimulatory effects of peptide A [309,310,314]. These studies suggest that peptide A isa key region of the II–III loop in RyR1 activation. However, additional experimentsin dysgenic myocytes showed that skeletal E–C coupling was not altered when thisregion was scrambled, deleted or replaced [302–304].

DHPR a1s regions other than the II–III loop may also be involved in skeletal-typeE–C coupling. The I–II loop was proposed to support weak E–C coupling [128], asynthetic C-terminal peptide (amino acids 1487–1506) was able to inhibit RyRs [320]and the recombinant III–IV loop competed with the II–III loop for binding to aRyR1 fragment [321]. Furthermore, expression of a1-subunit chimaeras in dysgenicmyocytes suggested that regions distinct from the II–III loop are involved in skeletal-type E–C coupling and the tetrad organization of DHPRs [322,323]. In addition, theb-subunit has recently been shown to interact directly with RyR1 and strengthenE–C coupling [324].

The type of E–C coupling depends not only on the DHPR isoform but also on theRyR isoform. Expression of RyR isoforms in skeletal myocytes derived from aknock-out mouse of RyR1 (also known as dyspedic mouse) demonstrated thatRyR1 restores skeletal-type E–C coupling, whereas expression of RyR2 or RyR3results in the cardiac type [325,326]. Mapping of the DHPR interaction site(s) onRyR1 has produced conflicting results. Screening of recombinant fragments encom-passing the large cytoplasmic N-terminal region of RyR1 identified residues922–1112 to be capable of binding the II–III loop of the DHPR a1s-subunit


[321,327]. The binding site was further localized to a 37-residue stretch (amino acids1076–1112) by use of RyR1/2 chimaeric fragments, and this region was also found tobind the a1s III–IV loop. However, the involvement of this RyR1 sequence inskeletal-type E–C coupling remains in question because expression of a chimaericRyR1, where this sequence was replaced with the equivalent RyR2 sequence,restored normal skeletal-type E–C coupling in dyspedic myocytes [328]. Furtherchimaeras between type 1 and type 2 RyRs identified two regions of RyR1, residues1635–2559 and 2659–3720, as essential for skeletal-type E–C coupling and formationof DHPR tetrads [328,329]. These results were also verified by RyR1/RyR3 chi-maeras and these studies were able to further refine the DHPR interaction sites toresidues 1924–2446 and 2644–3223 [330].

The use of RyR1/RyR3 chimaeras also suggested that a large N-terminal region(amino acids 1–1680) might participate in skeletal-type E–C coupling [331]. Deletionof divergent domain D2 within the RyR1N-terminus abolished skeletal-type E–Ccoupling; however, chimaeras between RyR1 and RyR2 or RyR3 indicated that thisdomain by itself is not sufficient to promote skeletal-type E–C coupling and mostprobably plays a structural role in RyR1 conformation [331,332]. Peptide A of thea1s DHPR subunit was reported to bind to two discrete RyR1 fragments, residues1021–1631 and 3201–3661, although deletion of only the second region abolishedpeptide A interaction with RyR1 [333]. Yeast two-hybrid analysis showed that thepeptide C region of the a1s-DHPR subunit interacted with RyR1 amino acids1837–2168, but expression of RyR1/2 chimaeras in dyspedic myocytes indicatedthat this region is only partially involved in skeletal-type E–C coupling [334].

Distinct RyR1 regions have also been implicated in DHPR interaction that do notinvolve the II–III loop of the a1s-subunit. The C-terminal tail of the a1s-subunit (orshorter peptides within this region) interacts with RyR1 residues 3609–3643 and 4064–4210 in a CaM-sensitive manner [291,335]. Also the DHPR b subunit binds to RyR1residues 3201–3661 through a cluster of positively charged amino acids [324]. Thus, allthe above studies taken together suggest that the DHPR-RyR association involvesmultiple regions on both proteins and/or is conformation sensitive.

9.2. FKBP

The cytosolic FKBP, FKBP12 and the closely related FKBP12.6 isoform are classi-fied as immunophilins as they are the receptor proteins for the immunosuppressantdrugs, FK506 and rapamycin, and are the smallest members of an extensive andubiquitous FKBP family (reviewed in [336]). Both isoforms have peptidyl-prolyl cis-trans isomerase activity that is inhibited by FK506 and rapamycin. Both FKBP12and FKBP12.6 consist of 108 amino acids including a cleaved N-terminal methionineresidue, and they share approximately 85% homology. The structures of both iso-forms, determined by X-ray crystallography, were found to be nearly identical toeach other and unaltered in their drug complexes [337–339]. Northern blot analysishas revealed transcripts for both isoforms in all mammalian tissues but significantlyhigher levels for FKBP12 [340].


RyR1 binds both FKBP12/12.6 with similar affinities, as shown byco-immunoprecipitation and co-purification assays and by GST-FKBP affinitychromatography [341–344]. However, native RyR1 from skeletal muscle is isolatedas a complex with FKBP12 because of relative abundance of FKBP12 over FKBP12.6.The RyR1-FKBP12 association in skeletal muscle is common to each of the five classesof vertebrates, i.e. mammals (rabbit), birds (chicken), reptiles (turtle), fish (salmon andrainbow trout) and amphibians (frog). The stoichiometry is four molecules of eitherFKBP12 or FKBP12.6 per RyR1 tetramer, i.e. one FKBP per RyR1 subunit.Mutation analysis identified three important residues (Q3, R18 and M49) in theFKBP12 interaction with RyR1 [345] and revealed that FKBP isomerase activity isnot required [342]. Surface plasmon resonance analysis of RyR1–FKBP12 bindingcharacteristics revealed a conformation-sensitive, high-affinity interaction; strongFKBP12 binding occurs when the channel is in its open state (Kd� 1nM), but it isfurther enhanced (by approximately 4–5 orders of magnitude) when the channel is inits closed configuration [346]. Using GST-FKBP12 affinity chromatography, theinteraction was found to be largely unaffected by temperature and pH changes, butit was ionic strength dependent (optimal at 100–200mM NaCl) [347]. FKBP12-boundRyR1 (in its closed state) has been visualized by cryo-electron microscopy and foundto be localized along the edge of the square-shaped cytoplasmic assembly, approxi-mately 12nm away from the putative channel pore (Fig. 1) [95].

RyR2 associates specifically with FKBP12.6 also with a stoichiometry of oneFKBP12.6 per RyR2 subunit [340,343,348,349]. The RyR2-FKBP12.6 associationin the heart is common in mammals and other vertebrates including chicken, fish andfrog. Although canine RyR2 binds FKBP12.6 exclusively, RyR2 from other speciesis also able to bind FKBP12 but with a 7- to 8-fold lower affinity, and thereforenative RyR2 from cardiac muscle is isolated as a complex with FKBP12.6. Mutationanalysis has identified three residues (Q31, N32 and F59) critical for selective bindingof FKBP12.6 to RyR2 [350] and revealed that enzymatic isomerase activity is notrequired [351]. The location of the FKBP12.6-binding site on RyR2 (in its open state)visualized by cryo-electron microscopy appears similar, although slightly displaced(by approximately 2 nm) from the FKBP12 site of RyR1 [352]. RyR3 is capable ofbinding both FKBP12/12.6 with similar affinities [353]. The immunosuppressantdrugs FK506 and rapamycin disrupt the RyR-FKBP interactions [343,354–356].

Initial reports using yeast two-hybrid assay or GST-FKBP affinity chromatogra-phy combined with limited RyR proteolysis have implicated the central RyR region(amino acids 2407–2520 in RyR1) in the interaction with FKBP12/12.6[234,353,357,358]. Furthermore, mutation analysis has identified V2461 in RyR1(and the corresponding V2322 in RyR3) as an important residue for FKBP bindingas assessed by co-immunoprecipitation, GST-FKBP-affinity chromatography andfunctional assays [353,359–361]. However, other studies, using very similar experi-mental approaches and RyRs with mutations or deletions in the central region, havequestioned the involvement of this domain in FKBP binding [273,360,362]. Twoalternative, very large domains have been proposed to form the FKBP12.6-bindingsite for the cardiac RyR isoform: the amino-terminus (residues 307–1855) [273,363]or the carboxyl-terminus (residues 3788–4967) [364]. The reason for the conflicting


results is unclear but may be due to isoform and/or species differences. Hence, theremay be multiple FKBP-binding sites in the RyR (at the N-terminus, central domainand C-terminus) with different binding properties and different sensitivities, depend-ing on channel state. Notably, the FKBP12-binding site visualized on RyR1 by cryo-electron microscopy at approximately 16 A resolution does not prove or disprove anyof the three proposed sites, because it appears to be in close proximity, but it does notoverlap with any of them [365].

Functional effects of FKBP12 on skeletal muscle RyR activity have been studiedin isolated channels or SR preparations following FKBP12 dissociation with FK506or rapamycin. SR Ca2+ flux measurements, single channel recordings andryanodine-binding assays have demonstrated that FKBP12 removal results inenhanced channel activation. In particular, FKBP12-deficient RyR1 channels havean increased sensitivity to Ca2+ or caffeine activation and reduced sensitivity tomillimolar Ca2+ or Mg2+ inhibition, whereas control values are restored uponreconstitution with FKBP12/12.6 [342,344,354,356,366–369]. Single channel record-ings of RyR1 have revealed pronounced subconductance activity upon FKBP12dissociation. FKBP12-deficient RyR1 channels, generated either by FK506 or rapa-mycin treatment (with or without subsequent removal of FKBP12-drug complexes),or recombinant channels expressed in insect cells devoid of FKBP12/12.6, or nativechannels isolated from FKBP12 knock-out mice, displayed openings to approxi-mately 25, 50 and 75% of the maximum conductance [356,366,367,370]; however,subconductance states are not always observed [368,369]. The effects of FKBP12.6on cardiac RyR are similar to that of FKBP12 on RyR1. Single channel recordings ofFKBP-deficient RyR2 channels, either by application of FK506 or rapamycin or bygene-targeted deletion of FKBP12 or FKBP12.6 in mice, also indicated subconduc-tance behaviour [355,370–372]. However, other studies found no effect on RyR2channel activity or conductance upon FKBP12.6 removal or addition [343,369].FKBP12/12.6 has also been proposed to enhance the functional coupling betweenadjacent RyR channels (termed ‘coupled gating’) in both skeletal and cardiac muscle[373,374], although FKBPs are not involved in their physical coupling [267]. Singlechannel recording studies have shown that addition of FKBP induced simultaneousopenings and closings of two channels incorporated into the planar lipid bilayer,whereas rapamycin treatment resulted in uncoupled channels. However, FKBPbinding occurs at the opposite site of the RyR–RyR contact region (which is closeto the D2 region) indicating that FKBP is not a direct intermediate protein but mayallosterically modulate tetramer–tetramer interactions [267].

FKBP12/12.6 functional effects have also been studied in isolated myocytes ormuscle fibres. FK506 and rapamycin-enhanced caffeine- and halothane-inducedskeletal muscle contractions [367,375] that are consistent with FKBP12-promotingchannel closure. These studies also suggested an involvement of FKBP12 dissocia-tion in MH (see Section 10.3.); however, MH responses were not detected in in vitrocaffeine contracture tests with FKBP12-deficient mice [376]. Depolarization-inducedCa2+ release studies revealed a role for an intact RyR1-FKBP12 association inskeletal-type E–C coupling. FKBP12-deficiency resulted in reduced depolarization-induced Ca2+ release and contraction [360,376–378]. As SR Ca2+ content was


normal, these findings suggest that FKBP12 dissociation suppresses the intrinsic abilityof the DHPR voltage sensors to efficiently activate Ca2+ release. Contrary to skeletalmuscle, FKBP12.6 deficiency in cardiomyocytes, induced by application of FK506 orrapamycin or by gene-targeted deletion of FKBP12.6, resulted in increased intracel-lular Ca2+ transients and contractions and also in increased Ca2+ spark frequencyand/or amplitude and duration [371,379,380]. The effects of FKBP12.6 have beeninvestigated in heterologous systems expressing recombinant RyR2 channels, whereco-expression of FKBP12.6 (but not FKBP12) suppressed spontaneous or agonist-induced Ca2+ release [381,382]. Adenovirus-mediated FKBP12.6 over-expression inisolated cardiomyocytes resulted in increased electrically evoked Ca2+ transients andSR Ca2+ content, whereas it decreased Ca2+ spark frequency, amplitude and duration[383–385]. Although increased Ca2+ transients upon FKBP12.6 over-expressionappears to contradict the results obtained by FK506 or rapamycin treatment (whereincreased Ca2+ transients were also observed), the former were due to increased SRCa2+ content, most likely because of suppressed RyR2 spontaneous activity.FKBP12.6 over-expression in transgenic mice resulted in enhanced contractility andcardiac output, although detailed analysis of potential adaptive changes was notcarried out [351]. These studies taken together indicate that FKBP12.6 has an inhibi-tory effect on RyR2 activity and promotes channel closure.

The physiological role of FKBP12/12.6 has been addressed by gene knock-outstudies in mice. FKBP12-deficient mice have normal skeletal muscle but severecardiac defects and die at embryonic stage or shortly after birth [370]. Mice with askeletal muscle-specific deletion of FKBP12 live normally, but certain types ofskeletal muscle have compromised contractile properties [376]. The generation ofmice deficient in FKBP12.6 by two independent groups has produced two differentphenotypes [372,380]. In both cases, mice had a normal phenotype; however, in onelaboratory mild cardiac hypertrophy was observed in male but not in female mice.This was discrepant with the second group, where both sexes presented with exercise-induced cardiac arrhythmias. The reason for disparate phenotypes is unclear, but itcould be due to different mouse strains.

In summary, FKBP12/12.6 stabilizes channel conductance, promotes channelclosure, supports coupled gating between adjacent channels and enhances skeletal-type E–C coupling. In contrast, FKBP dissociation results in enhanced sensitivity ofCa2+ activation and ‘leaky’ channels. Because of their profound effects on RyRchannel properties, FKBPs are considered essential components of the RyR complexin both skeletal and cardiac muscle. It should be noted that defective regulation ofthe RyR2-FKBP12.6 association has been implicated in the pathogenesis of heartfailure and cardiac arrhythmias (see Sections 10.1. and 10.2.).

9.3. CaM

CaM is a ubiquitous, highly conserved 17-kDa protein that has a high affinity forCa2+ and participates in numerous Ca2+ signalling processes (reviewed in [386]).CaM has a dumbbell-shaped structure consisting of N-terminal and C-terminal


globular domains connected by a flexible linker, each containing two EF-hand Ca2+-binding motifs. CaM binds all three mammalian RyRs with nanomolar affinityirrespective of whether it is in a Ca2+-free or Ca2+-bound state, although affinityis slightly higher at micromolar calcium [387–391]. There are four CaMmolecules perRyR channel, i.e. one CaM per RyR subunit, at both low and high Ca2+ levels. This1:1 stoichiometry replaces earlier findings based on chemically labelled CaM (either[125I] or fluorescent label) or mapping studies with RyR fragments, suggestingmultiple binding sites for CaM per RyR subunit [392–395]. The RyR1–CaM bindingsite has been visualized by cryo-electron microscopy; CaM was localized at theRyR1 periphery, approximately 10 nm away from the putative channel pore(Fig. 1) [95].

The primary binding site for both Ca2+-free and Ca2+-bound CaM lies withinresidues 3614–3643 of RyR1 from assessment of protease protection assays,cysteine modification protection assays, peptide-binding analysis and mutationanalysis [226,387,396,397]. CaM is suggested to bind at a site of intersubunitcontact and protects RyR1 from oxidation, with identified RyR1 residues, W3620and L3624, being critical for the interaction. The presence of Ca2+ shifts theposition of CaM several amino acids N-terminal of the Ca2+-free site, because asmaller C-terminal-truncated peptide (amino acids 3614–3634) retained the inter-action with the Ca2+-bound, but not Ca2+-free, CaM [396]. Further studies indi-cate that the CaM C-terminal globular domain binds RyR1 at 3614–3643, whereasthe N-terminal domain binds to an adjacent RyR1 subunit at residues 1975–1999[294,398]. The primary CaM site is also conserved in RyR2 and RyR3, becausemutations or deletion of RyR2 residues 3583–3603 or RyR3 residues 3469–3489abolished CaM binding [391,399]. Mutation analysis also suggests that additional,isoform-specific regions are involved in the differential modulation of the threeRyRs by CaM [391,400].

CaM has a biphasic, Ca2+-dependent effect on RyR1 activity from SR Ca2+

flux measurements, single channel recordings and ryanodine binding assays[388,393,401–404]. At low nanomolar Ca2+ levels, CaM activates RyR1, but athigher Ca2+ concentrations (�1 mM), it inhibits the channel. However, CaM is notessential for E–C coupling, because mutation of its binding site on the RyR1 didnot affect depolarization-induced Ca2+ release [405]. Similar to RyR1, RyR3activity is also stimulated by CaM at low Ca2+ levels but inhibited at high Ca2+

[25,391,406]. In contrast, RyR2 is inhibited by CaM at all Ca2+ concentrations,although inhibition is more pronounced at high Ca2+ [170,390,402,404]. CaM hasbeen speculated to be involved in Ca2+ release termination, as it has an inhibitoryeffect for all RyRs at micromolar Ca2+. However, CaM was found to associateand dissociate from RyR2 on a time scale of seconds to minutes [390], whichsuggests that CaM occupancy of the channel remains unchanged during thecardiac cycle, and the kinetics of CaM-regulated RyR2 activity suggested only afacilitatory role in Ca2+ release termination [407]. In skeletal muscle, CaM wasshown to increase Ca2+ spark frequency, whereas other spark properties wereunaffected, suggesting a role for CaM in initiation, but not termination, of Ca2+

release [408].


CaM may also affect SR Ca2+ release by acting on other proteins including theDHPR Ca2+ channel, CaMKII and calcineurin, a CaM-activated protein phospha-tase (reviewed in [409]).

9.4. Sorcin

Sorcin is a 22-kDa member of the penta-EF-hand Ca2+ binding protein family,which binds Ca2+ with high affinity (reviewed in [410]). It has a widespreadexpression including the heart, where it localizes along the Z-lines, in particular thedyadic junctions of T-tubules and SR TC [411–413]. Sorcin is implicated inmodulation of cardiac E–C coupling, because it associates with RyR2 andDHPR channels [411,414] and translocates from cytosol to membrane-embeddedtargets upon increasing intracellular Ca2+ (to approximately 200mM) [413]. Sorcininteracts with DHPR through the cytoplasmic C-terminal tail of the a1-subunit [414],whereas the sorcin-binding site on RyR2 remains to be determined. Single channelrecordings and ryanodine-binding assays have demonstrated isoform-specificeffects; submicromolar sorcin levels result in near-complete RyR2 inhibition that isCa2+-independent,whereas it has a stimulatory effect on RyR1 [415]. Kineticsanalysis indicates that RyR2 inhibition occurs within milliseconds, suggesting thatsorcin may be involved in termination of Ca2+ release [413].

Sorcin studies in vivo and in isolated cardiomyocytes have yielded disparateresults. Evidence of a role for sorcin in suppression of E–C coupling consistentwith RyR2 inhibition is provided by some studies but not all. Sorcin depressedintracellular Ca2+ transients and contraction and decreased Ca2+ sparkfrequency, when dialysed into cells through a patch-pipette, following adenovirus-mediated over-expression, in transgenic mice with cardiac-specific over-expression[412,413,416]. However, there were also some differences. In isolated rabbitcardiomyocytes infected with an adenoviral sorcin construct, depressed Ca2+

transients were attributed to a reduced SR Ca2+ content resulting from enhancedactivity of the Na+–Ca2+ exchanger, although decreased Ca2+ spark frequency wasindeed due to RyR2 inhibition. In transgenic animals, sorcin over-expressionaccelerated the rate of L-type Ca2+ current inactivation, whereas L-type Ca2+

current density and kinetics were unaffected in the other two studies. Moresignificant differences were reported by three other studies where an enhancementof E–C coupling upon sorcin over-expression was observed [417–419]. Adenoviraltransfer of sorcin in isolated cardiomyocytes or in vivo in mouse and rat heartsresulted in enhanced Ca2+ transients and contractility. One study also reportedstimulation of SR Ca2+ uptake activity by sorcin and enhanced SR Ca2+ contentas the underlying mechanism [419]. The reason(s) for the above discrepanciesremain unclear. It could be due to species differences, although most of thestudies used rat cardiomyocytes, and/or due to different sorcin levels. The abovestudies also suggest that sorcin has other targets in addition to the RyR and DHPRchannels, although a direct interaction has not been demonstrated for theNa+–Ca2+exchanger or the SR Ca2+ pump. Sorcin is also known to form dimers


and tetramers, a process that is Ca2+ and H+ dependent [420], and it is possible thatits target binding is differentially affected by its oligomeric state.

9.5. CSQ, triadin and junctin

CSQ is a highly acidic glycoprotein that comprises the main Ca2+ storage protein inthe SR lumen. Isolated first from striated muscle, where it is primarily expressed,CSQ is a low-affinity, high-capacity Ca2+-binding protein (approximately 40molCa2+ per mol protein) that undergoes conformational changes upon binding Ca2+

[421,422]. CSQ forms dimers, tetramers and eventually polymers as the Ca2+ con-centration increases [423]. CSQ polymers are visible in electron micrographs ofjunctional muscle preparations, as electron-dense material exclusively in the TC ofthe SR, and they appear to be anchored near the luminal face of RyRs [2].

Two isoforms of CSQ are known, comprising approximately 390 residues andsharing 86% homology, termed the skeletal and cardiac type due to their relativeabundance in skeletal and cardiac muscle, respectively [424,425]. Their sequencesreveal no TM domains and no apparent SR retention signal. CSQs are rich in acidicresidues (approximately 30%), most of which are involved in Ca2+ binding at theC-terminal region, with pairs of acidic residues creating a net surface charge to whichCa2+ binds [423]. Ca2+ causes conformational changes in the monomer and inducesCSQ polymerisaton [423]. Micromolar Ca2+ (10mM) causes compaction of the CSQmonomer, whereas further increases (10–100mM) lead to dimerization and thenformation of linear polymers at millimolar Ca2+. Hence, CSQ exists as a stablepolymer at physiological free luminal Ca2+ concentration of approximately 1mM[426]. Both skeletal and cardiac CSQs bind more Ca2+ ions than predicted by theirnet negative charges (–80 and –69, respectively), with approximately 50 and 36mol ofCa2+ per protein molecule, respectively, at �5mM Ca2+, as they form their poly-meric states [427]. Skeletal and cardiac CSQ have almost identical three-dimensionalstructures revealed by X-ray crystallography, each with three similar domains and atopology reminiscent of Escherichia coli thioredoxin [423,427].

CSQ is anchored to the junctional face of the SR membrane through interactionswith two SR integral proteins, triadin and junctin, and together with the RyR, thefour proteins form a macromolecular complex [428,429]. Junctin and triadin bindingare mediated through an aspartate-rich region (amino acids 354–367) at the C-terminus of CSQ [430]. This region is also implicated in Ca2+ binding, suggestingthat Ca2+ and triadin/junctin compete for binding to the same site. CSQ can bedissociated from the junctional face membrane by either low (<10mM) or high Ca2+

concentrations (10mM) or high ionic strength [429,431]. The interaction betweenCSQ and RyR is either direct (although the two main RyR luminal loops do not bindCSQ [432]) or indirect through triadin and junctin [429]. Recent evidence suggeststhat CSQ is dissociated from RyR at Ca2+ concentrations >4mM [191].

Triadin was isolated from skeletal muscle as a 95-kDa membrane glycoproteinfrom junctional SR [433]. It is now known to be encoded by a single gene but to existin multiple forms in both skeletal and cardiac muscle because of alternative splicing


[434–438]. Four subtypes have been found in skeletal muscle, termed Trisk 95, 51, 49and 32 based on their molecular weights, with the first two being the predominantones. In the heart, three subtypes have been identified, named CT1, 2 and 3 withmolecular weights of 32, 35 and 75 kDa, with CT1 being the predominant one. Allthe isoforms share a common N-terminal cytoplasmic segment, a single TM domain(residues 47–67) and luminal parts of variable length, with unique sequences at theC-terminal end. A single TM segment is the accepted membrane topology based onhydropathy plot analysis and immunolocalization studies [434,439]. Triadin formsoligomers through disulphide bridges [440]. Interaction of triadin with RyR ismediated through both cytoplasmic and luminal domains [428,441,442]. Triadinbinding to the SR membrane cytoplasmic face involves residues 18–46 and is Ca2+

dependent, with an optimum at <10mM. Binding to the SR luminal side involves atriadin sequence of alternating positive and negative residues (amino acids 210–224,known as the KEKE motif) interacting with the most C-terminal RyR luminal loop(M8–M10 loop) where three negatively charged residues (D4878, D4907 and E4908)are important for the interaction. Triadin interaction with CSQ is also mediatedthrough the KEKE motif [443]. Interestingly, Trisk 49 and 32 do not bind RyR,although they contain both the N-terminal cytoplasmic domain and the KEKE motif[436].

Junctin, a protein of 26 kDa encoded by a single gene, is also an integral mem-brane protein localized in the junctional SR of both skeletal and cardiac muscle [444].It has a similar predicted structure to triadin with a short, 21-residue cytoplasmicN-terminus, a single TM domain with the highly charged remainder residing in theSR lumen. Unlike triadin, junctin is not glycosylated, and it does not form disul-phide-linked oligomers. Through its luminal domain, junctin binds RyR and triadinin a Ca2+-independent manner, whereas binding to CSQ is disrupted by either low-or high Ca2+ concentrations [429].

It is generally assumed that the primary role of triadin and junctin is to dock CSQto the RyR, though their individual roles remain to be investigated in detail. In singlechannel recordings, triadin was reported to inhibit RyR1 but only when added to thecytoplasmic side of the channel [445], whereas luminal addition of triadin and junctinto purified RyR2 led to activation [190]. These differences could be due to the use ofnative versus purified RyRs (see below). The functions of triadin have been studied intransgenic animals with cardiac-specific over-expression [446,447]. Transgenic ani-mals developed cardiac hypertrophy, had altered muscle contraction and SR Ca2+

handling and enhanced SR Ca2+ content. Over-expression of triadin (CT1) wasaccompanied by reduced expression of junctin and RyR2, although the number offunctional RyR2s was unchanged. Similar results were obtained with transgenicanimals over-expressing junctin, although SR Ca2+ content was reduced [448,449].Although these studies suggest that triadin and junctin play an important role in theCa2+ release process, interpretation of the results is complicated by the variousadaptive changes in these animals. To overcome these problems, the role of triadinwas studied following acute, adenovirus-mediated over-expression in primary myo-cytes. Over-expression of CT1 in cardiomyocytes enhanced E–C coupling and Ca2+

spark frequency and increased the activity of RyR2 channels suggesting a


stimulatory role for triadin [450]. The role of triadin in skeletal muscle is not clear,because acute over-expression of Trisk 95 (but not Trisk 51) in skeletal musclemyotubes did not affect pharmacological activation of RyR1 or Ca2+-inducedCa2+ release but abolished depolarization-induced Ca2+ release, i.e. skeletal-typeE–C coupling [451].

Reports of functional effects of CSQ on RyR have been contradictory. It appearshowever, that the disparate results are due to the use of native or purified channelsthat lack the complement of accessory proteins, in particular triadin and junctin[431]. Initial single channel recording studies using either purified channels or chan-nels, where the association with triadin and/or junctin is likely to be disrupted,reported that CSQ addition activates the RyR [431,445,452,453]. However, studieswith native channels or channels, where the macromolecular complex with triadinand junctin was reconstituted, demonstrated that CSQ inhibits both skeletal andcardiac RyRs [190,191,431]. The latter studies also revealed the need for an intactmacromolecular complex for luminal Ca2+ regulation of the RyR (see Section 7.2.).

An inhibitory effect of CSQ on RyR activity is supported by CSQ over-expressingtransgenic mice, because these animals had higher SR Ca2+ content, but Ca2+

transients and Ca2+ spark frequency were reduced [454–456]. Notably, the mutantmice developed severe hypertrophy and heart failure and/or had altered expression ofRyR, triadin and junctin complicating the interpretation of results. Acute, adeno-virus-mediated over-expression in primary myocytes produced conflicting results.Limited over-expression of CSQ (approximately 1.6-fold) in cardiomyocytes resultedin increased SR Ca2+ content but reduced E–C coupling, in agreement with aninhibitory role for CSQ [457]. In contrast, a higher over-expression of CSQ (approxi-mately 4-fold) resulted in increased E–C coupling in cardiomyocytes [458], as well asin skeletal muscle cells [459]. Thus, it seems that CSQ functional effects depend on thestoichiometry of association with the RyR/triadin/junctin/CSQ macromolecularcomplex.

The importance of CSQ in control of SR Ca2+ release was demonstrated by theidentification of homozygous mutations in patients with CPVT (see Section 10.2.).Functional characterization of two mutations (R33Q and D307H) revealed twodistinct mechanisms of action, although they both resulted in arrhythmogenicmembrane potentials following b-adrenergic stimulation in cardiomyocytes over-expressing the mutant proteins [460,461]. Adenovirus-mediated over-expression ofR33Q CSQ enhanced E–C coupling and Ca2+ spark frequency despite normal(total) SR Ca2+ content (versus wild type) [461]. In single channel recordings, theR33Q CSQ mutation abolished its inhibitory effect on RyR2, and it was thereforeproposed that this mutation results in enhanced RyR2 channel sensitivity to lumi-nal Ca2+ activation. In contrast, over-expression of D307H CSQ resulted indecreased E–C coupling and Ca2+ spark frequency consistent with the concomitantreduction in SR Ca2+ store content [460]. In vitro characterization of this mutantrevealed that it had a profoundly altered conformation and reduced affinity forCa2+, triadin and junctin [462]. It is unclear how the two CSQ mutations result incardiac arrhythmias while they produce opposing effects on RyR2-mediated SRCa2+ release.


10. RyR pathophysiology

10.1. Heart failure

Heart failure resulting from different forms of cardiomyopathy is defined as theinability of the heart to pump sufficient blood to meet the body’s metabolic demands(reviewed in [463]). It is a leading cause of disability and sudden death because ofventricular arrhythmias. In the early stages of heart failure, compensatory neurohor-monal mechanisms are triggered, including the sympathetic nervous system, andcatecholamine-induced stimulation of b-adrenergic receptors leads to production ofcAMP and activation of PKA. PKA-mediated phosphorylation enhances L-type Ca2+

current, whereas phosphorylation of phospholamban relieves its inhibition of the SRCa2+ ATPase pump, resulting in higher SR Ca2+ content. The net effect is enhance-ment of E–C coupling and improvement of cardiac function. However, in later stagesof heart failure, calcium homeostasis is impaired, and contractile dysfunction is causedprimarily by reduced intracellular Ca2+ transients and by concomitant decrease in theSR Ca2+ content (reviewed in [464]). There is now considerable evidence that diastolicSR Ca2+ leak, through RyR2, contributes to reduced SR Ca2+ content and systolicCa2+ transients in heart failure [227,234,259,465,466]. However, there is controversywith regard to the mechanism that leads to RyR2-mediated Ca2+ leak.

An extensive series of studies from one laboratory have provided compellingevidence that in heart failure, chronic b-adrenergic stimulation results in RyR2‘hyper-phosphorylation’ at S2808 by PKA, inducing dissociation of the stabilizingFKBP12.6 co-protein, thereby enhancing channel activity [234,241,245,246,467–471].This hypothesis was based on the following observations: (i) failing cardiomyocytesdisplayed higher phosphorylation levels exclusively at S2808, (ii) S2808 was uniquelyand exclusively phosphorylated by PKA, whereas CaMKII phosphorylated a differ-ent, unique residue, S2814, (iii) RyR2 hyper-phosphorylation was due to decreasedlocal protein phosphatase and cAMP hydrolysis activities (reduced levels of PP1,PP2A and phosphodiesterase 4D3 associated with RyR2), (iv) FKBP12.6 levelsassociated with RyR2 were severely reduced in failing cardiomyocytes, PKA-treatedchannels or S2808D mutant channels (mimicking constitutive phosphorylation) butwere normal in S2808A channels (that cannot be phosphorylated), (v) RyR2 chan-nels isolated from failing cardiomyocytes or treated with PKA or S2808D (but notS2808A) mutant channels displayed enhanced channel activity and subconductancebehaviour characteristic of FKBP12.6 dissociation and consistent with SR Ca2+ leakin cells, (vi) b-adrenergic receptor blockers reversed RyR2 hyper-phosphorylation,FKBP12.6 dissociation and channel activation in heart failure, (vii) RyR2 S2808Atransgenic mice were resistant to heart failure, (viii) a benzothiazepine, JTV519, wasfound to restore the RyR2-FKBP12.6 association in failing cardiomyocytes, PKA-treated channels or S2808D mutant channels and (ix) JTV519 protected mice fromheart failure, but it was without effect in FKBP12.6-deficient animals.

An independent group has supported some of the above findings, includingincreased RyR2 phosphorylation, FKBP12.6 dissociation and the beneficial effects


of beta-blockers and JTV519 in heart failure [472–475]. However, it also reportedsome important differences, namely that the initial critical step leading to RyR2dysregulation is a defective interdomain interaction that is induced by oxidativestress [227,281]. These conclusions are based on the finding that disruption of theRyR N-terminal–central domain interaction by the DPc10 peptide or by oxidizingreagents resulted in Ca2+ leak through RyR2 without removing FKBP12.6 althoughit facilitated PKA-mediated phosphorylation and FKBP12.6 dissociation. Impor-tantly, RyR2 was shown to be oxidized, and the interdomain interaction was dis-rupted in an animal model of heart failure but was restored by antioxidantadministration. In isolated failing cardiomyocytes, JTV519 also restored the inter-domain interaction and enhanced Ca2+ transients without reversing RyR2 phos-phorylation state or reassociation of FKBP12.6. These results suggest that oxidativestress-induced disruption of interdomain interactions within RyR2 is the primarycause of Ca2+ leak, whereas phosphorylation and FKBP12.6 dissociation are theconsequence, rather than the cause, of RyR2 leaky channels.

In addition, certain aspects of the hyper-phosphorylation hypothesis have beenquestioned by results from several other laboratories that used very similar experi-mental approaches. First, S2808 was found to be highly phosphorylated in normalhearts even at rest, and to be a substrate for CaMKII and PKG in addition to PKA,and that it is not a unique PKA site (see Section 7.6.). Second, PKA-mediated RyR2phosphorylation levels and channel activity were similar in failing and normalcardiomyocytes [242,476], whereas the increased S2808 phosphorylation that wasfound in some animal models of heart failure was unrelated to b-adrenergic stimula-tion and RyR2 dysfunction [244,477]. In line with these studies, PKA activation inpermeabilized cardiomyocytes obtained from phospholamban knock-out mice didnot affect Ca2+ spark frequency (under conditions that excluded PKA effects onDHPR Ca2+ current and SR Ca2+ content) [258,259]. Third, no cardiomyopathiesor heart failures were reported in FKBP12.6 knock-out mice (although mildhypertrophy for male mice was reported in one of the two models) [372,380]. Fourth,PKA-dependent phosphorylation or S2808D substitution did not cause FKBP12.6dissociation, and S2808D mutant channel activity was identical to wild type[242,247,363,382,476]. A recent study detected a reduced association between RyR2and FKBP12.6 (by approximately 40%) in failing cardiomyocytes but also reported adecrease in cellular FKBP12.6 levels, which at the mRNA level was reduced byapproximately 50% [478]. This finding suggests that the reduced RyR2-FKBP12.6association often seen in heart failure could simply be a result of lower FKBP12.6expression levels.

An alternative mechanism in the pathogenesis of heart failure involves CaMKII-dependent phosphorylation of RyR2. It has been shown that CaMKII activity isincreased upon sustained b-adrenergic stimulation and in heart failure [478–481]. Inan animal model of heart failure, RyR2 had increased phosphorylation levels due tohigher levels of associated CaMKII, and failing cardiomyocytes had an increasedRyR2-mediated SR Ca2+ leak that was inhibited by CaMKII inhibitors [478].Chronic over-expression of CaMKIIdC in transgenic mice resulted in contractiledysfunction and heart failure, with decreased Ca2+ transients and SR Ca2+ content,


as well as changes in the expression profile of proteins involved in E–C coupling[261,479]. Although these adaptive changes make it difficult to pinpoint the primarydefect, there was increased CaMKII-dependent phosphorylation of RyR2 that wasassociated with increased Ca2+ spark frequency. In contrast, transgenic mice over-expressing a CaMKII inhibitory peptide were resistant to maladaptive remodellingassociated with heart failure and had preserved contractility and Ca2+ homeostasis[481]. It should be noted that CaMKII-mediated phosphorylation of RyR2 did notresult in FKBP12.6 dissociation [259,262].

Increased SR Ca2+ leak may also be due to increased luminal Ca2+ activation ofRyR2. It has been reported in an animal model of heart failure that there was anincrease in Ca2+ spark frequency despite lower SR Ca2+ content, and the RyR2studied by single channel recordings was found to have an enhanced sensitivity toluminal Ca2+ activation [466].

10.2. Arrhythmogenic cardiac diseases

CPVT is an inherited arrhythmogenic disease characterized by adrenergicallymediated bidirectional or polymorphic ventricular tachycardia leading to syncopeand/or sudden cardiac death (reviewed in [482,483]). Although it is relatively rare,CPVT is a highly malignant disease (mortality rates of 30–35%) with incompletepenetrance, manifesting in childhood and adolescence. Patients with CPVT havestructurally normal hearts and typically present with ventricular arrhythmias becauseof physical or emotional stress, and they can also be induced reproducibly by theinfusion of catecholamines. Administration of b-adrenergic receptor blockers is thestandard treatment; however, it is not always effective. CPVT has been linked to twocardiac SR proteins, RyR2 and CSQ. Mutations in RyR2 cause a dominant form ofCPVT, whereas mutations in CSQ are linked to an autosomal recessive form. Muta-tions in RyR2 are also associated with arrhythmogenic right ventricular dysplasiatype 2 (ARVD2). Patients with ARVD2 exhibit progressive fibro-fatty replacementof the right ventricular myocardium, in addition to stress-induced polymorphicventricular tachycardia. However, the structural abnormalities are relatively mildand ARVD2mimics the CPVT phenotype, suggesting an overlap between the twodiseases. To date, 68 point mutations and a 2-residue insertion have been identified inRyR2 that tend to segregate in three distinct regions: the N-terminus, the centraldomain and the C-terminus containing the membrane-spanning pore-forming helices(Fig. 5).

Several studies have characterized some RyR2mutations following expression ofrecombinant channels in cardiac (HL-1) or other (HEK293) cell lines [372,484–491].Most studies suggest that mutations do not affect channel state in resting conditions,although enhanced basal activity for at least some mutants (N4104K, R4497C1 andN4895D) has been reported [484,486]. All studies indicate that RyR2mutationsresult in increased SR Ca2+ release after channel activation; however, more detailedexamination to reveal the molecular mechanism(s) underlying channel dysfunctionhas produced contradictory results. In a cell-based assay, some mutant channels were


found to have altered sensitivity to caffeine or cytoplasmic Ca2+ activation, whereasother mutants (R176Q/T2504M and L433P) had markedly reduced sensitivity tocytoplasmic Ca2+ inactivation [487,489]. The latter observation is in line withreduced Mg2+ inhibition of other mutants (P2328S, Q4201R and V4653F) whenstudied by single channel recordings [488]. It has also been proposed that PKA-mediated phosphorylation, with the ensuing FKBP12.6 dissociation, results inabnormal SR Ca2+ release during b-adrenergic stimulation, in common with themechanism proposed in heart failure [372,470,488,492]. This proposal was based onthe observations that mutant RyR2 channels (S2246L, P2328S, R2474S, Q4201R,R4497C and V4653F) had reduced affinity for FKBP12.6, mice deficient (orhaploinsufficient) in FKBP12.6 presented with exercise-induced ventricular arrhyth-mias similar to CPVT patients and JTV519 prevented triggered arrhythmias byrestoring the RyR2-FKBP12.6 association. However, FKBP12.6 dissociation upon

E1724K

S2246L P2328SN2386I

V186M

R2474ST2504M

A2403T

V2475F

Y2392C

A2394GR2359QF2331S

E2311DL3778F

C3800F

S3938R

G3946S

E4076K

S4124T

T4196A

F4020L

N4097S

Q4201R

R4497C

A4607PT4158P

E4146K

V4653F

G4662SG4671R

H4762P

R4959QE4950K

N4895DV4880AI4867MA4860G

F4851CI4848V

V4771I

A4556T

F4499C

M4504IA4510T

L2534V

L433P

P466A

R420W

I419FR176Q

R169QP164S

A77V

R2401H/L

R414L/C

A2387P/T

P4902L/S

Ins EY4657/8H4108N/Q

N4104K/I

A2254V

V2306I

Fig. 5. Ryanodine receptor 2 (RyR2) mutations associated with arrhythmogenic diseases. Schematic to

illustrate catecholaminergic polymorphic ventricular tachycardia/arrhythmogenic right ventricular dyspla-

sia type 2 (CPVT/ARVD2) mutations (red circles) clustered primarily in three discrete regions of the RyR2

polypeptide: the N-terminal (residues 77–466), the central (residues 2246–2534) and the C-terminal domain

(residues 3778–4959). Image kindly prepared by Dr. N. Lowri Thomas (Wales Heart Research Institute,

Cardiff University, UK) (See Color Plate 37, p. 534).


PKA-dependent phosphorylation has been heavily contested, and JTV519 wasshown to act independently of FKBP12.6 (see Section 10.1.). In addition, otherstudies found normal FKBP12.6 interaction with channels harbouring the samemutations [485,490,493], whereas b-blockers or JTV519 could not prevent cardiacarrhythmias in a mouse knock-in RyR2mutant model for R4497C [493,494].

An alternative hypothesis proposes that RyR2mutations exclusively enhance thesensitivity of the channel to luminal Ca2+ activation [486,490]. It was found thatspontaneous Ca2+ release occurred at a lower Ca2+ store threshold in cells expres-sing mutant channels (R176Q/T2504M, L433P, S2246L, R2474S, N4104K, Q4201R,R4497C, I4867M and N4895D), and single channel recordings indicated enhancedluminal Ca2+ activation but unaltered cytoplasmic Ca2+ sensitivity. This hypothesisis consistent with involvement of CSQ mutations in CPVT (see Section 9.5.).Enhanced sensitivity to luminal calcium is likely to involve long range, conforma-tional changes and allosteric interactions, at least for the N-terminal and centraldomain mutations. The disruption of regulatory interactions between the N-terminusand central domain by CPVT-linked mutations has been proposed to underlie thedefective regulation of RyR2mutant channels (see Section 8.2.). In addition, RyR2mutations (N4104K and R4497C) were shown by a protein fragment complementa-tion assay to result in altered interactions within a region (amino acids 3722–4610)suggested to transduce cytoplasmic signals to the pore-forming TM assembly [491].

The generation of two mouse knock-in models of R4497C and R176Q linked toCPVT and ARVD2, respectively, has yielded important information about themechanism of inherited, RyR2-linked, triggered arrhythmias [493–495]. These ani-mal models demonstrated that RyR2mutations pre-dispose the mouse heart to thedevelopment of ventricular arrhythmias upon exercise or infusion of b-adrenergicagonists. About 30% of R4497C heterozygous mice developed ventriculartachycardia, which was unrelated to the relative expression levels of the wild-typeand mutant alleles [494], consistent with incomplete penetrance of the CPVT disease,where some family members carrying RyR2mutations are asymptomatic.Electrophysiological analysis indicated that triggered arrhythmias are mediated bydelayed after-depolarizations [493–495], in common with cardiac arrhythmias inheart failure (reviewed in [496,497]). The likely mechanism is that RyR2-mediateddiastolic SR Ca2+ leak activates the plasma membrane Na+–Ca2+ exchanger (whichexchanges one Ca2+ for three Na+) thereby generating a net inward current thatcould induce diastolic depolarizations (called delayed after-depolarizations). Delayedafter-depolarizations could in turn elicit pre-mature action potentials and initiationof arrhythmias. Interestingly, R4497C knock-in mice occasionally demonstrateddelayed after-depolarizations at rest, which were enhanced upon b-adrenergicstimulation, suggesting that mutant RyR2 has a basal defect that is exacerbatedby b-adrenergic stimulation [493]. This phenotype is consistent with the clinicalobservations that b-blockers provide only partial protection in CPVT patients.Mice heterozygous for R176Q exhibited catecholamine-induced ventricular tachy-cardia and had structurally normal hearts (although right ventricular end-diastolicvolume was decreased), suggesting that ARVD2 and CPVT may represent subtle butdistinct variations of a single disorder [495].


The use of animal knock-in models should help to identify the molecular mechan-ism(s) underlying RyR2 dysfunction. In vitro functional assays have suggestedseveral mechanisms including altered sensitivity to luminal Ca2+ activation, cyto-plasmic Ca2+ and Mg2+ inhibition, inter-domain interactions, phosphorylationstatus and FKBP12.6 interaction. The reason for the disparate results is not clear.It is notable that expression of recombinant channels in HEK293 cells results inhomotetramers, whereas in HL-1 (which contain endogenous wild type RyR2), thisresults in heterotetramers, mimicking the heterozygous CPVT (or ARVD2) patients.HL-1 cells also have an appropriate genetic background, expressing the complementof cardiac-specific accessory proteins. However, different findings were reported forthe same RyR2mutation, even when expressed in the same cell line. At present, theimportant unresolved issue is to determine the primary causative defect. For exam-ple, altered inter-domain interactions within the RyR2 structure may be the primarycause, resulting in a channel that is more prone to open and/or remain open. Thiscould be manifested as enhanced sensitivity to cytoplasmic and/or luminal Ca2+

activation and/or reduced sensitivity to Ca2+ and Mg2+ inhibition. However, giventhe dispersion of mutations across the RyR2 sequence, it may be unlikely that there isa common mechanism to account for channel dysregulation.

10.3. Neuromuscular disorders

Three clinically distinct hereditary skeletal muscle disorders are known to be asso-ciated with mutations in the gene encoding RyR1: MH, central core disease (CCD)and multi-minicore disease (MmD) (reviewed in [498]). MH is an autosomal domi-nant pharmacogenetic disorder of skeletal muscle, triggered in pre-disposed indivi-duals by inhalation of anaesthetics (e.g. halothane) or by depolarising musclerelaxants (e.g. succinylcholine). Susceptible individuals respond with skeletal musclerigidity, hypermetabolism, tachycardia, unstable and rising blood pressure and even-tually dramatic hyperthermia. MH is a life-threatening disease and is treated with theRyR1 antagonist dantrolene.

CCD is a congenital myopathy characterized by hypotonia during infancy, prox-imal muscle weakness, delayed motor development and reduced muscle bulk. It isusually inherited as an autosomal-dominant disease although recessive forms havealso been described. Diagnosis of CCD is determined through histological identifica-tion of large amorphous areas (cores) that lack mitochondria and thus oxidativeenzyme activity and cover a considerable area of primarily type 1muscle fibres.MmD is an autosomal recessive congenital myopathy that is characterized histolo-gically by the presence of multiple cores devoid of mitochondria and with disorga-nized sarcomeric structures. Multiple small cores can occur in both type 1 and 2fibres that do not run the entire length of the muscle fibre. Patients affected by MmDhave symptoms similar to CCD, and there may be some overlap between the twodiseases. More than 80 different mutations in the RyR1 gene have so far beenassociated with MH, CCD and MmD. They are mostly point mutations althoughsmall, in-frame deletions or out-of-frame insertions have also been found. RyR1


mutations cluster in three separate regions: in the N-terminus, the central domainand the C-terminus containing the channel pore, with very few mutations outsidethese regions. These three regions correspond to the ones identified in RyR2-associated arrhythmogenic diseases (see Section 10.2.), which might suggest acommon mechanism of action. RyR1 is the only gene so far linked to CCD, whereasmutations in RyR1 account for the majority of MH cases and only a minority ofMmD cases. Although both MH- and CCD-associated mutations are found in allthree regions, MH-linked mutations are mostly in the N-terminal and central regions,whereas CCD-linked mutations predominate in the C-terminus. Interestingly, someRyR1mutations cause both MH and CCD.

The functional effects of MH-/CCD-linked mutations have been extensively stu-died by a combination of methods. These include single-channel recordings andryanodine-binding assays of isolated channels, as well as Ca2+ release measurementsin heterologous (HEK293 and COS-7) cell lines, or in primary dyspedic or normalmyocytes expressing recombinant mutant RyR1, or of endogenous mutant channelsin skeletal myocytes or lymphocytes [isolated from affected individuals, pigs with anaturally occurring MH mutation (R614C1) or a mouse knock-in MH model,Y522S]. These studies have indicated that RyR1mutations in the N-terminal andcentral regions and most mutations in the C-terminus result in higher sensitivity topharmacological agonists such as caffeine and halothane, as well as to activation byplasma membrane depolarization, and also in reduced sensitivity to inactivation byMg2+ and Ca2+ [499–510]. Although differences exist between studies, it was foundthat RyR1mutant channels had enhanced basal activity at rest but to a variableextent, resulting in higher resting intracellular Ca2+ concentration and lower Ca2+

store content and consequently reduced magnitude of Ca2+ release [505,507,510–516]. CCD-linked mutations had the most profound effect on calcium store depletionirrespective of their location on the RyR1 sequence, whereas MH-linked mutationshad smaller or no effects. Contradictory findings have been reported for a subset ofC-terminal CCD-associated mutations located within the pore-forming luminal loop(e.g. I4898T1). Although some studies have shown that these mutations also produce‘leaky’ channels with increased resting intracellular Ca2+ concentration and lowerCa2+ store content [512,516–518], others found normal cytoplasmic and SR Ca2+

levels but severely reduced sensitivity to Ca2+ activation and depolarization- oragonist-induced Ca2+ release [513,519,520].

The above investigations suggest that although both MH- and CCD-linked muta-tions result in ‘hyper-sensitive’ channels and increased basal activity, only CCD-linkedmutations cause a substantial Ca2+ leak from the SR that cannot be compensated forby increased Ca2+ uptake. This leads to a partially depleted SR Ca2+ store and in turndecreased Ca2+ release during E–C coupling and therefore muscle weakness. Alterna-tively, compromised E–C coupling is due to RyR1mutations within the pore-formingsequences that result in reduced Ca2+ permeability. In MH, RyR1mutations result inreduced inactivation of channels by Ca2+ andMg2+ and therefore prolonged elevationin intracellular Ca2+ concentration and muscle rigidity. Disruption of inter-domaininteractions involved in stabilization of the channel in a closed state has also beenimplicated in MH (see Section 8.2.).


11. Conclusions

Since its purification and molecular cloning in the late 1980s, the RyR has beenextensively studied using a variety of approaches that range from single channel analysisto animal models. These studies have contributed to our understanding of the RyRproperties at the molecular level and also in the context of the intact cell and wholeanimal. We now know a lot about the biophysical properties of the channel, itsregulation by endogenous and exogenous effectors and its physiological mechanismof activation in skeletal and cardiac muscle. There are, however, aspects that we still donot understand fully. Of paramount importance is the lack of information about thethree-dimensional structure of the protein and of mechanism(s) that satisfactorilyexplain termination of SR Ca2+ release despite the inherent positive feedback of CICR.

Defective RyR2 regulation, either induced (in heart failure) or inherited (in CPVTand ARVD2), is directly involved in cardiac disease and sudden death. Extensiveresearch is currently focused on the delineation of the underlying pathway(s) ofRyR2-mediated pathology, but thus far, conflicting reports have only produced aconfusing picture. Clearly much more research is needed in order to clarify this fieldand identify the causative molecular defect(s). It is only then that with the combinedefforts of basic scientists and clinicians, treatments can be developed aimed atrestoring RyR2 function and therefore cellular Ca2+ homeostasis and cardiacoutput.

Acknowledgements

We are grateful to Dr. Zheng Liu, Dr. Leon D’Cruz and Dr. N. Lowri Thomas fortheir kind assistance in preparing the images used in Fig. 1, 3 and 5, respectively. Ourresearch is funded by grants from the British Heart Foundation, The WellcomeTrust, European Union (CONTICA programme) and Medical Research Council.

Note

1. Numbering given for the human sequence.

References

1. Berridge, M., Lipp, P. and Bootman, M. (2000) Nat Rev Mol Cell Biol 1, 11–21.

2. Franzini-Armstrong, C. (1999) FASEB J 13, S266–S270.

3. Sutko, J., Airey, J., Welch, W. and Ruest, L. (1997) Pharmacol Rev 49, 53–98.

4. Fleischer, S., Ogunbunm, I., Dixon, M. and Fleer, E. (1985) Proc Natl Acad Sci USA 82, 7256–7259.

5. Pessah, I., Francini, A., Scales, D., Waterhouse, A. and Casida, J. (1986) J Biol Chem 261, 8643–8648.

6. Campbell, K., Knudson, C., Imagawa, T., Leung, A., Sutko, J., Kahl, S., Raab, C. and Madson, L.

(1987) J Biol Chem 262, 6460–6463.

7. Inui, M., Saito, A. and Fleischer, S. (1987) J Biol Chem 262, 1740–1747.


8. Lai, F., Erickson, H., Rousseau, E., Liu, Q. and Meissner, G. (1988) Nature 331, 315–319.

9. Inui, M., Saito, A. and Fleischer, S. (1987) J Biol Chem 262, 15637–15642.

10. Anderson, K., Lai, F., Liu, Q., Rousseau, E., Erickson, H. and Meissner, G. (1989) J Biol Chem 264,

1329–1335.

11. Lai, F., Misra, M., Xu, L., Smith, H. and Meissner, G. (1989) J Biol Chem 264, 16776–16785.

12. Imagawa, T., Smith, J., Coronado, R. and Campbell, K. (1987) J Biol Chem 262, 16636–16643.

13. Hymel, L., Inui, M., Fleischer, S. and Schindler, H. (1988) Proc Natl Acad Sci USA 85, 441–445.

14. Sutko, J. and Airey, J. (1996) Physiol Rev 76, 1027–1071.

15. Franzini-Armstrong, C. and Protasi, F. (1997) Physiol Rev 77, 699–729.

16. Takeshima, H., Nishimura, S., Matsumoto, T., Ishida, H., Kangawa, K., Minamino, N., Matsuo, H.,

Ueda, M., Hanaoka, M., Hirose, T. and Numa, S. (1989) Nature 339, 439–445.

17. Zorzato, F., Fujii, J., Otsu, K., Phillips, M., Green, N., Lai, F., Meissner, G. and MacLennan, D.

(1990) J Biol Chem 265, 2244–2256.

18. Nakai, J., Imagawa, T., Hakamata, Y., Shigekawa, M., Takeshima, H. and Numa, S. (1990) FEBS

Lett 271, 169–177.

19. Otsu, K., Willard, H., Khanna, V., Zorzato, F., Green, N. and MacLennan, D. (1990) J Biol Chem

265, 13472–13483.

20. Hakamata, Y., Nakai, J., Takeshima, H. and Imoto, K. (1992) FEBS Lett 312, 229–235.

21. Chen, S., Vaughan, D., Airey, J., Coronado, R. and MacLennan, D. (1993) Biochemistry 32,

3743–3753.

22. Chen, S., Leong, P., Imredy, J., Bartlett, C., Zhang, L. and MacLennan, D. (1997) Biophys J 73,

1904–1912.

23. Imagawa, T., Nakai, J., Takeshima, H., Nakasaki, Y. and Shigekawa, M. (1992) J Biochemistry 112,

508–513.

24. Bhat, M., Hayek, S., Zhao, J., Zang, W., Takeshima, H., Wier, W. and Ma, J. (1999) Biophys J 77,

808–816.

25. Chen, S., Li, X., Ebisawa, K. and Zhang, L. (1997) J Biol Chem 272, 24234–24246.

26. Saeki, K., Obi, I., Ogiku, N., Hakamata, Y. and Matsumoto, T. (1998) Life Sci 63, 575–588.

27. Sorrentino, V. and Volpe, P. (1993) Trends Pharmacol Sci 14, 98–103.

28. Airey, J., Beck, C., Murakami, K., Tanksley, S., Deerinck, T., Ellisman, M. and Sutko, J. (1990) J

Biol Chem 265, 14187–14194.

29. Olivares, E., Tanksley, S., Airey, J., Beck, C., Ouyang, Y., Deerinck, T., Ellisman, M. and Sutko, J.

(1991) Biophys J 59, 1153–1163.

30. Lai, F., Liu, Q., Xu, L., El-Hashem, A., Kramarcy, N., Sealock, R. and Meissner, G. (1992) Am J

Physiol 263, C365–C372.

31. Murayama, T. and Ogawa, Y. (1992) J Biochem 112, 514–522.

32. O’Brien, J., Meissner, G. and Block, B. (1993) Biophys J 65, 2418–2427.

33. Airey, J., Grinsell, M., Jones, L., Sutko, J. and Witcher, D. (1993) Biochemistry 32, 5739–5745.

34. Oyamada, H., Murayama, T., Takagi, T., Iino, M., Iwabe, N., Miyata, T., Ogawa, Y. and Endo, M.

(1994) J Biol Chem 269, 17206–17214.

35. Ottini, L., Marziali, G., Conti, A., Charlesworth, A. and Sorrentino, V. (1996) Biochem J 315,

207–216.

36. Seok, J., Xu, L., Kramarcy, N., Sealock, R. and Meissner, G. (1992) J Biol Chem 267, 15893–15901.

37. Takeshima, H., Nishi, M., Iwabe, N., Miyata, T., Hosoya, T., Masai, I. and Hotta, Y. (1994) FEBS

Lett 337, 81–87.

38. Sakube, Y., Ando, H. and Kagawa, H. (1993) Ann NY Acad Sci 707, 540–545.

39. Tunwell, R., Wickenden, C., Bertrand, B., Shevchenko, V., Walsh, M., Allen, P. and Lai, F. (1996)

Biochem J 318, 477–487.

40. Zorzato, F., Sacchetto, R. and Margreth, A. (1994) Biochem Biophys Res Commun 203, 1725–1730.

41. Futatsugi, A., Kuwajima, G. and Mikoshiba, K. (1995) Biochem J 305, 373–378.


42. Marziali, G., Rossi, D., Giannini, G., Charlesworth, A. and Sorrentino, V. (1996) FEBS Lett 394,

76–82.

43. Miyatake, R., Furukawa, A., Masayuki, M., Iwahashi, K., Nakamura, K., Ichikawa, Y. and Suwaki,

H. (1996) FEBS Lett 395, 123–126.

44. Jiang, D., Xiao, B., Li, X. and Chen, S. (2003) J Biol Chem 278, 4763–4769.

45. Kimura, T., Nakamori, M., Lueck, J., Pouliquin, P., Aoike, F., Fujimura, H., Dirksen, R.,

Takahashi, M., Dulhunty, A. and Sakoda, S. (2005) Hum Mol Genet 14, 2189–2200.

46. Xiao, B., Masumiya, H., Jiang, D., Wang, R., Sei, Y., Zhang, L., Murayama, T., Ogawa, Y., Lai, F.,

Wagenknecht, T. and Chen, S. (2002) J Biol Chem 277, 41778–41785.

47. Beutner, G., Sharma, V., Giovannucci, D., Yule, D. and Sheu, S. (2001) J Biol Chem 276,

21482–21488.

48. Mitchell, K., Pinton, P., Varadi, A., Tacchetti, C., Ainscow, E., Pozzan, T., Rizzuto, R. and Rutter,

G. (2001) J Cell Biol 155, 41–51.

49. Kuwajima, G., Futatsugi, A., Niinobe, M., Nakanishi, N. and Mikoshiba, K. (1992) Neuron 9,

1133–1142.

50. Lai, F., Dent, M., Wickenden, C., Xu, L., Kumari, G., Misra, M., Lee, H., Sar, M. and Meissner, G.

(1992) Biochem J 288, 553–564.

51. Ouyang, Y., Deerinck, T., Walton, P., Airey, J., Sutko, J. and Ellisman, M. (1993) Brain Res 620,

269–280.

52. Ledbetter, M., Preiner, J., Louis, C. and Mickelson, J. (1994) J Biol Chem 269, 31544–31551.

53. Furuichi, T., Furutama, D., Hakamata, Y., Nakai, J., Takeshima, H. and Mikoshiba, K. (1994)

J Neurosci 14, 4794–4805.

54. Giannini, G., Conti, A., Mammarella, S., Scrobogna, S. and Sorrentino, V. (1995) J Cell Biol 128,

893–904.

55. Neylon, C., Richards, S., Larsen, M., Agrotis, A. and Bobik, A. (1995) Biochem Biophys Res Commun

215, 814–821.

56. Martin, C., Chapman, K., Seckl, J. and Ashley, R. (1998) Neuroscience 85, 205–216.

57. Murayama, T. and Ogawa, Y. (1997) J Biol Chem 272, 24030–24037.

58. Jeyakumar, L., Copello, J., O’Malley, A., Wu, G.-M., Grassucci, R., Wagenknecht, T. and Fleischer, S.

(1998) J Biol Chem 273, 16011–16020.

59. Takeshima, H., Iino, M., Takekura, H., Nishi, M., Kuno, J., Minowa, O., Takano, H. and Noda, T.

(1994) Nature 369, 556–559.

60. Takeshima, H., Komazaki, S., Hirose, K., Nishi, M., Noda, T. and Iino, M. (1998) EMBO J 17,

3309–3316.

61. Takeshima, H., Ikemoto, T., Nishi, M., Nishiyama, N., Shimuta, M., Sugitani, Y., Kuno, J., Saito, I.,

Saito, H., Endo, M., Iino, M. and Noda, T. (1996) J Biol Chem 271, 19649–19652.

62. Bertocchini, F., Ovitt, C., Conti, A., Barone, V., Scholer, H., Bottinelli, R., Reggiani, C. and

Sorrentino, V. (1997) EMBO J 16, 6956–6963.

63. Balschun, D., Wolfer, D., Bertocchini, F., Barone, V., Conti, A., Zuschratter, W., Missiaen, L., Lipp, H.,

Frey, U. and Sorrentino, V. (1999) EMBO J 18, 5264–5273.

64. Futatsugi, A., Kato, K., Ogura, H., Li, S., Nagata, E., Kuwajuma, G., Tanaka, K., Itohara, S. and

Mikoshiba, K. (1999) Neuron 24, 701–713.

65. Williams, A., West, D. and Sitsapesan, R. (2001) Q Rev Biophys 34, 61–104.

66. Smith, J., Imagawa, T., Ma, J., Fill, M., Campbell, K. and Coronado, R. (1988) J Gen Physiol 92,

1–26.

67. Liu, Q., Lai, F., Rousseau, E., Jones, R. and Meissner, G. (1989) Biophys J 55, 415–424.

68. Lindsay, A., Manning, S. and Williams, A. (1991) J Physiol 439, 463–480.

69. Tinker, A. and Williams, A. (1992) J Gen Physiol 100, 479–493.

70. Murayama, T., Oba, T., Katayama, E., Oyamada, H., Oguchi, K., Kobayashi, M., Otsuka, K. and

Ogawa, Y. (1999) J Biol Chem 274, 17297–17308.

71. Kasai, M., Kawasaki, T. and Yamamoto, K. (1992) J Biochem 112, 197–203.


72. Tinker, A. and Williams, A. (1993) J Gen Physiol 102, 1107–1129.

73. Tu, Q., Velez, P., Brodwick, M. and Fill, M. (1994) Biophys J 67, 2280–2285.

74. Tinker, A. and Williams, A. (1995) Biophys J 68, 111–120.

75. Tinker, A., Lindsay, A. and Williams, A. (1992) J Membr Biol 127, 149–159.

76. Tu, Q., Velez, P., Cortes-Gutierrez, M. and Fill, M. (1994) J Gen Physiol 103, 853–867.

77. Tinker, A. and Williams, A. (1993) Biophys J 65, 852–864.

78. Xu, L., Jones, R. and Meissner, G. (1993) J Gen Physiol 101, 207–233.

79. Mead, F., Sullivan, D. and Williams, A. (1998) J Membr Biol 163, 225–234.

80. Xu, L., Tripathy, A., Pasek, D. and Meissner, G. (1999) J Biol Chem 274, 32680–32691.

81. Mead, F. and Williams, A. (2004) Biophys J 87, 3814–3825.

82. Welch, W., Rheault, S., West, D. and Williams, A. (2004) Biophys J 87, 2335–2351.

83. Marks, A. (1996) Physiol Rev 76, 631–649.

84. Anyatonwu, G., Buck, E. and Ehrlich, B. (2003) J Biol Chem 278, 45528–45538.

85. Zhao, M., Li, P., Li, X., Zhang, L., Winkfein, R. and Chen, S. (1999) J Biol Chem 274, 25971–25974.

86. Radermacher,M., Rao, V., Grassucci, R., Frank, J., Timerman, A., Fleischer, S. andWagenknecht, T.

(1994) J Cell Biol 127, 411–423.

87. Serysheva, I., Orlova, E., Chiu, W., Sherman, M., Hamilton, S. and Van Heel, M. (1995) Nat Struct

Biol 2, 18–24.

88. Sharma, M., Penczek, P., Grassucci, R., Xin, H., Fleischer, S. and Wagenknecht, T. (1998) J Biol

Chem 273, 18429–18434.

89. Serysheva, I., Hamilton, S., Chiu, W. and Ludtke, S. (2005) J Mol Biol 345, 427–431.

90. Samso, M., Wagenknecht, T. and Allen, P. (2005) Nat Struct Mol Biol 12, 539–544.

91. Ludtke, S., Serysheva, I., Hamilton, S. and Chiu, W. (2005) Structure 13, 1203–1211.

92. Orlova, E., Serysheva, I., van Heel, M., Hamilton, S. and Chiu, W. (1996) Nat Struct Biol 3,

547–552.

93. Serysheva, I., Schatz, M., Van Heel, M., Chiu, W. and Hamilton, S. (1999) Biophys J 77, 1936–1944.

94. Sharma, M., Jeyakumar, L., Fleischer, S. and Wagenknecht, T. (2000) J Biol Chem 275, 9485–9491.

95. Wagenknecht, T., Radermacher, M., Grassucci, R., Berkowitz, J., Xin, H. and Fleisher, S. (1997)

J Biol Chem 272, 32463–32471.

96. Liu, Z., Zhang, J., Sharma, M., Li, P., Chen, S. and Wagenknecht, T. (2001) Proc Natl Acad Sci

USA 98, 6104–6109.

97. Baker, M., Serysheva, I., Sencer, S., Wu, Y., Ludtke, S., Jiang, W., Hamilton, S. and Chiu, W.

(2002) Proc Natl Acad Sci USA 99, 12155–12160.

98. Liu, Z., Wang, R., Zhang, J., Chen, S. and Wagenknecht, T. (2005) J Biol Chem 280, 37941–37947.

99. Benacquista, B., Sharma, M., Samso, M., Zorzato, F., Treves, S. and Wagenknecht, T. (2000)

Biophys J 78, 1349–1358.

100. Liu, Z., Zhang, J., Li, P., Chen, S. and Wagenknecht, T. (2002) J Biol Chem 277, 46712–46719.

101. Zhang, J., Liu, Z., Masumiya, H., Wang, R., Jiang, D., Li, F., Wagenknecht, T. and Chen, S. (2003)

J Biol Chem 278, 14211–14218.

102. Liu, Z., Zhang, J., Wang, R., Chen, S. and Wagenknecht, T. (2004) J Mol Biol 338, 533–545.

103. Wang, J., Needleman, D., Seryshev, A., Aghdasi, B., Slavik, K., Liu, S.-Q., Pedersen, S. and

Hamilton, S. (1996) J Biol Chem 271, 8387–8393.

104. Bhat, M., Zhao, J., Takeshima, H. and Ma, J. (1997) Biophys J 73, 1329–1336.

105. Bhat, M., Zhao, J., Zang, W., Balke, C., Takeshima, H., Wier, W. and Ma, J. (1997) J Gen Physiol

110, 749–762.

106. Xu, X., Bhat, M., Nishi, M., Takeshima, H. and Ma, J. (2000) Biophys J 78, 1270–1281.

107. George, C., Jundi, H., Thomas, N., Scoote, M., Walters, N., Williams, A. and Lai, F. (2004) Mol

Biol Cell 15, 2627–2638.

108. Marty, I., Villaz, M., Arlaud, G., Bally, I. and Ronjat, M. (1994) Biochem J 298, 743–749.

109. Grunwald, R. and Meissner, G. (1995) J Biol Chem 270, 11338–11347.


110. Du, G., Sandhu, B., Khanna, V., Guo, X. and MacLennan, D. (2002) Proc Natl Acad Sci USA 99,

16725–16730.

111. Du, G., Avila, G., Sharma, P., Khanna, V., Dirksen, R. and MacLennan, D. (2004) J Biol Chem

279, 37566–37574.

112. Doyle, D., Morais Cabral, J., Pfuetzner, R., Kuo, A., Gulbis, J., Cohen, S., Chait, B. and

MacKinnon, R. (1998) Science 280, 69–77.

113. Jiang, Y., Lee, A., Chen, J., Cadene, M., Chait, B. and MacKinnon, R. (2002) Nature 417, 523–526.

114. Balshaw, D., Gao, L. and Meissner, G. (1999) Proc Natl Acad Sci USA 96, 3345–3347.

115. Gao, L., Balshaw, D., Xu, L., Tripathy, A., Xin, C. and Meissner, G. (2000) Biophys J 79, 828–840.

116. Du, G., Guo, X., Khanna, V. and MacLennan, D. (2001) J Biol Chem 276, 31760–31771.

117. Chen, S., Li, P., Zhao, M., Li, X. and Zhang, L. (2002) Biophys J 82, 2436–2447.

118. Wang, R., Zhang, L., Bolstad, J., Diao, N., Brown, C., Ruest, L., Welch, W., Williams, A. and

Chen, S. (2003) J Biol Chem 278, 51557–51565.

119. Wang, R., Bolstad, J., Kong, H., Zhang, L., Brown, C. and Chen, S. (2004) J Biol Chem 279,

3635–3642.

120. Du, G. and MacLennan, D. (1998) J Biol Chem 273, 31867–31872.

121. Wang, Y., Xu, L., Pasek, D., Gillespie, D. and Meissner, G. (2005) Biophys J 89, 256–265.

122. Xu, L., Wang, Y., Gillespie, D. and Meissner, G. (2006) Biophys J 90, 443–453.

123. Ferguson, D., Schwartz, H. and Franini-Armstrong, C. (1984) J Cell Biol 99, 1735–1742.

124. Saito, A., Inui, M., Radermacher, M., Frank, J. and Fleischer, S. (1988) J Cell Biol 107, 211–219.

125. Schneider, M. and Chandler, W. (1973) Nature 242, 244–246.

126. Rios, E. and Brum, G. (1987) Nature 325, 717–720.

127. Tanabe, T., Beam, K., Powell, J. and Numa, S. (1988) Nature 336, 134–139.

128. Tanabe, T., Beam, K., Adams, B. and Niidome, T. (1990) Nature 346, 567–569.

129. Jacquemond, V., Csernoch, L., Klein, M. and Schneider, M. (1991) Biophys J 60, 867–873.

130. Marty, I., Robert, M., Villaz, M., De Jongh, K., Lai, Y., Catterall, W. and Ronjat, M. (1994) Proc

Natl Acad Sci USA 91, 2270–2274.

131. Block, B., Leung, A., Campbell, K. and Franzini-Armstrong, C. (1988) J Cell Biol 107, 2587–2600.

132. Takekura, H., Bennett, L., Tanabe, T., Beam, K. and Franzini-Armstrong, C. (1994) Biophys J 67,

793–804.

133. Franzini-Armstrong, C. and Kish, J. (1995) J Muscle Res Cell Motil 16, 319–324.

134. Nakai, J., Dirksen, R., Nguyen, H., Pessah, I., Beam, K. and Allen, P. (1996) Nature 380, 72–75.

135. Sun, X., Protasi, F., Takahashi, M., Takeshima, H., Ferguson, D. and Franzini-Armstrong, C.

(1995) J Cell Biol 129, 659–667.

136. Carl, S., Felix, K., Caswell, A., Brandt, N., Ball, W., Vaghy, P., Meissner, G. and Ferguson, D.

(1995) J Cell Biol 129, 672–682.

137. Protasi, F., Sun, X. and Franzini-Armstrong, C. (1996) Dev Biol 173, 265–278.

138. Reuter, H. and Beeler, G.J. (1969) Science 163, 399–401.

139. Tanabe, T., Mikami, A., Numa, S. and Beam, K. (1990) Nature 344, 451–453.

140. Cleemann, L. and Morad, M. (1991) J Physiol 432, 283–312.

141. Sham, J., Cleeman, L. and Morad, M. (1995) Proc Natl Acad Sci USA 92, 121–125.

142. Gomez, A., Valdivia, H., Cheng, H., Lederer, M., Santana, L., Cannel, M., McCune, S., Altschuld,

R. and Lederer, W. (1997) Science 276, 800–806.

143. Gomez, A., Guatimosim, S., Dilly, K., Vassort, G. and Lederer, W. (2001) Circulation 104, 688–693.

144. He, J.-Q., Conklin, M., Foell, J., Wolff, M., Haworth, R., Coronado, R. and Kamp, T. (2001)

Cardiovasc Res 49, 298–307.

145. Balijepalli, R., Lokuta, A., Maertz, N., Buck, J., Haworth, R., Valdivia, H. and Kamp, T. (2003)


146. Song, L.-S., Sobie, E., McCulle, S., Lederer, W., Balke, C. and Cheng, H. (2006) Proc Natl Acad Sci

USA 103, 4305–4310.


147. Fill, M. and Copello, J. (2002) Physiol Rev 82, 893–922.

148. Stern, M. and Cheng, H. (2004) Cell Calcium 35, 591–601.

149. Zucchi, R. and Ronca-Testoni, S. (1997) Pharmacol Rev 49, 1–51.

150. Rousseau, E., Smith, J. and Meissner, G. (1987) Am J Physiol 253, C364–C368.

151. Buck, E., Zimanyi, I., Abramson, J. and Pessah, I. (1992) J Biol Chem 267, 23560–23567.

152. Zimanyi, I., Buck, E., Abramson, J., Mack, M. and Pessah, I. (1992)Mol Pharmacol 42, 1049–1057.

153. Fessenden, J., Chen, L., Wang, Y., Paolini, C., Franzini-Armstrong, C., Allen, P. and Pessah, I.

(2001) Proc Natl Acad Sci USA 98, 2865–2870.

154. Masumiya, H., Li, P., Zhang, L. and Chen, S. (2001) J Biol Chem 276, 39727–39735.

155. Du, G., Guo, X., Khanna, V. and MacLennan, D. (2001) Proc Natl Acad Sci USA 98, 13625–13630.

156. Chiu, A., Diaz-Munoz, M., Hawkes, M., Brush, K. and Hamilton, S. (1990) Mol Pharmacol 37,

735–741.

157. Pessah, I. and Zimanyi, I. (1991) Mol Pharmacol 39, 679–689.

158. Wang, J., Needleman, D. and Hamilton, S. (1993) J Biol Chem 268, 20974–20982.

159. Holmberg, S. and Williams, A. (1990) Biochim Biophys Acta 1022, 187–193.

160. Callaway, C., Seryshev, A., Wang, J., Slavik, K., Needleman, D., Cantu, C., Wu, Y., Jayaraman, T.,

Marks, A. and Hamilton, S. (1994) J Biol Chem 269, 15876–15884.

161. Witcher, D., McPherson, P., Kahl, S., Lewis, T., Bentley, P., Mullinnix, M., Windass, J. and

Campell, K. (1994) J Biol Chem 269, 13076–13079.

162. Tanna, B., Welch, W., Ruest, L., Sutko, J. and Williams, A. (2000) J Gen Physiol 116, 1–9.

163. Tanna, B., Welch, W., Ruest, L., Sutko, J. and Williams, A. (2003) J Gen Physiol 121, 551–561.

164. Meissner, G., Darling, E. and Eveleth, J. (1986) Biochemistry 25, 236–244.

165. Smith, J., Coronado, R. and Meissner, G. (1986) J Gen Physiol 88, 573–588.

166. Pessah, I., Stambuk, R. and Casida, J. (1987) Mol Pharmacol 31, 232–238.

167. Bull, R. and Marengo, J. (1993) FEBS Lett 331, 223–227.

168. Percival, A., Williams, A., Kenyon, J., Grinsell, M., Airey, J. and Sutko, J. (1994) Biophys J 67,

1834–1850.

169. Meissner, G., Rios, E., Tripathy, A. and Pasek, D. (1997) J Biol Chem 272, 1628–1638.

170. Meissner, G. and Henderson, J. (1987) J Biol Chem 262, 3065–3073.

171. Ashley, R. and Williams, A. (1990) J Gen Physiol 95, 981–1005.

172. Chu, A., Fill, M., Stefani, E. and Entman, M. (1993) J Membr Biol 135, 49–59.

173. Sitsapesan, R. and Williams, A. (1994) Biophys J 67, 1484–1494.

174. Laver, D., Roden, L., Ahern, G., Eager, K., Junankar, P. and Dulhunty, A. (1995) J Membr Biol

147, 7–22.

175. Schiefer, A., Meissner, G. and Isenberg, G. (1995) J Physiol 489, 337–348.

176. Sonnleitner, A., Conti, A., Bertocchini, F., Schindler, H. and Sorrentino, V. (1998) EMBO J 17,

2790–2798.

177. Ma, J. (1995) Biophys J 68, 893–899.

178. Copello, J., Barg, S., Onoue, H. and Fleiscer, S. (1997) Biophys J 73, 141–156.

179. Marengo, J., Hidalgo, C. and Bull, R. (1998) Biophys J 74, 1263–1277.

180. Li, P. and Chen, S. (2001) J Gen Physiol 118, 33–44.

181. Donoso, P., Prieto, H. and Hidalgo, C. (1995) Biophys J 68, 507–515.

182. Lamb, G., Cellini, M. and Stephenson, D. (2001) J Physiol 531, 715–728.

183. Sitsapesan, R. and Williams, A. (1994) J Membr Biol 137, 215–226.

184. Sitsapesan, R. and Williams, A. (1995) J Membr Biol 146, 133–144.

185. Tripathy, A. and Meissner, G. (1996) Biophys J 70, 2600–2615.

186. Xu, L. and Meissner, G. (1998) Biophys J 75, 2302–2312.

187. Gyorke, I. and Gyorke, S. (1998) Biophys J 75, 2801–2810.

188. Laver, D., O’Neill, E. and Lamb, G. (2004) J Gen Physiol 124, 741–758.


189. Ching, L., Williams, A. and Sitsapesan, R. (2000) Circ Res 87, 201–206.

190. Gyorke, I., Hester, N., Jones, L. and Gyorke, S. (2004) Biophys J 86, 2121–2128.

191. Beard, N., Casarotto, M., Wei, L., Varsanyi, M., Laver, D. and Dulhunty, A. (2005) Biophys J 88,

3444–3454.

192. Lamb, G. and Stephenson, D. (1994) J Physiol 478, 331–339.

193. Laver, D., Baynes, T. and Dulhunty, A. (1997) J Membr Biol 156, 213–229.

194. Copello, J., Barg, S., Sonnleitner, A., Porta, M., Diaz-Sylvester, P., Fill, M., Schindler, H. and

Fleischer, S. (2002) J Membr Biol 187, 51–64.

195. Xu, L., Mann, G. and Meissner, G. (1996) Circ Res 79, 1100–1109.

196. Liu, W., Pasek, D. and Meissner, G. (1998) Am J Physiol 274, C120–C128.

197. Meissner, G. (1984) J Biol Chem 259, 2365–2374.

198. Laver, D., Lenz, G. and Lamb, G. (2001) J Physiol 537, 763–778.

199. Kermode, H., Williams, A. and Sitsapesan, R. (1998) Biophys J 74, 1296–1304.

200. Manunta, M., Rossi, D., Simeoni, I., Butelli, E., Romanin, C., Sorrentino, V. and Schindler, H.

(2000) FEBS Lett 471, 256–260.

201. Abramson, J., Zable, A., Favero, T. and Salama, G. (1995) J Biol Chem 270, 29644–29647.

202. Aghdasi, B., Zhang, J.-Z., Wu, Y., Reid, M. and Hamilton, S. (1997) J Biol Chem 272, 3739–3748.

203. Eager, K., Roden, L. and Dulhunty, A. (1997) Am J Physiol 272, C1908–C1918.

204. Zable, A., Favero, T. and Abramson, J. (1997) J Biol Chem 272, 7069–7077.

205. Xia, R., Stangler, T. and Abramson, J. (2000) J Biol Chem 275, 36556–36561.

206. Boraso, A. and Williams, A. (1994) Am J Physiol 267, H1010–H1016.

207. Favero, T., Zable, A. and Abramson, J. (1995) J Biol Chem 270, 25557–25563.

208. Kawakami, M. and Okabe, E. (1998) Mol Pharmacol 53, 497–503.

209. Zima, A., Copello, J. and Blatter, L. (2004) J Physiol 555, 727–741.

210. Xia, R., Webb, J., Gnall, L., Cutler, K. and Abramson, J. (2003) Am J Physiol 285, C215–C221.

211. Cherednichenko, G., Zima, A., Feng, W., Schaefer, S., Blatter, L. and Pessah, I. (2004) Circ Res 94,

478–486.

212. Sanchez, G., Pedroso, Z., Domenech, R., Hidalgo, C. and Donoso, P. (2005) J Mol Cell Cardiol 39,

982–991.

213. Meszaros, L., Minarovic, I. and Zahradnikova, A. (1996) FEBS Lett 380, 49–52.

214. Stoyanovsky, D., Murphy, T., Anno, P., Kim, Y.-M. and Salama, G. (1997) Cell Calcium 21, 19–29.

215. Xu, L., Eu, J., Meissner, G. and Stamler, J. (1998) Science 279, 234–237.

216. Eu, J., Sun, J., Xu, L., Stamler, J. and Meissner, G. (2000) Cell 102, 499–509.

217. Aracena, P., Sanchez, G., Donoso, P., Hamilton, S. and Hidalgo, C. (2003) J Biol Chem 278,

42927–42935.

218. Cheong, E., Tumbev, V., Stoyanovsky, D. and Salama, G. (2005) Cell Calcium 38, 481–488.

219. Aracena, P., Tang, W., Hamilton, S. and Hidalgo, C. (2005) Antioxid Redox Signal 7, 870–881.

220. Sun, J., Xu, L., Eu, J., Stamler, J. and Meissner, G. (2001) J Biol Chem 276, 15625–15630.

221. Oba, T., Murayama, T. and Ogawa, Y. (2002) Am J Physiol 282, C684–C692.

222. Bull, R., Marengo, J., Finkelstein, J., Behrens, M. and Alvarez, O. (2003) Am J Physiol 285,

C119–C128.

223. Donoso, P., Aracena, P. and Hidalgo, C. (2000) Biophys J 79, 279–286.

224. Wu, Y., Aghdasi, B., Dou, S., Zhang, J., Liu, S. and Hamilton, S. (1997) J Biol Chem 272,

25051–25061.

225. Liu, G., Abramson, J., Zable, A. and Pessah, I. (1994) Mol Pharmacol 45, 189–200.

226. Zhang, J., Wu, Y., Williams, B., Rodney, G., Mandel, F., Strasburg, G.M. and Hamilton, S. (1999)

Am J Physiol 276, C46–C53.

227. Yano, M., Okuda, S., Oda, T., Tokuhisa, T., Tateishi, H., Mochizuki, M., Noma, T., Doi, M.,

Kobayashi, S., Yamamoto, T., Ikeda, Y., Ohkusa, T., Ikemoto, N. and Matsuzaki, M. (2005)

Circulation 112, 3633–3643.


228. Takasago, T., Imagawa, T. and Shikegawa, M. (1989) J Biochem 106, 872–877.

229. Chu, A., Sumbilla, C., Inesi, G., Jay, S. and Campbell, K. (1990) Biochemistry 29, 5899–5905.

230. Takasago, T., Imagawa, T., Furukawa, K., Ogurusu, T. and Shikegawa, M. (1991) J Biochem 109,

163–170.

231. Witcher, D., Kovacs, R., Schulman, H., Cefali, D. and Jones, L. (1991) J Biol Chem 266,

11144–11152.

232. Hohenegger, M. and Suko, J. (1993) Biochem J 296, 303–308.

233. Suko, J., Maurer-Fogy, I., Plank, B., Bertel, O., Wyskowsky, W., Hohenegger, M. and Hellmann,

G. (1993) Biochim Biophys Acta 1175, 193–206.

234. Marx, S., Reiken, S., Hisamatsu, Y., Jayaraman, T., Burkhoff, D., Rosemblit, N. and Marks, A.

(2000) Cell 101, 365–376.

235. Marx, S., Reiken, S., Hisamatsu, Y., Gaburjakova, M., Gaburjakova, J., Yang, Y.-M., Rosemblit,

N. and Marks, A. (2001) J Cell Biol 153, 699–708.

236. Currie, S., Loughrey, C., Craig, M.-A. and Smith, G. (2004) Biochem J 377, 357–366.

237. Hain, J., Nath, S., Mayrleitner, M., Fleischer, S. and Schindler, H. (1994) Biophys J 67, 1823–1833.

238. Hain, J., Onoue, H., Mayrleitner, M., Fleischer, S. and Schindler, H. (1995) J Biol Chem 270,

2074–2081.

239. Dulhunty, A., Laver, D., Curtis, S., Pace, S., Haarmann, C. and Gallant, E. (2001) Biophys J 81,

3240–3252.

240. Rodriguez, P., Bhogal, M. and Colyer, J. (2003) J Biol Chem 278, 38593–38600.

241. Wehrens, X., Lehnart, S., Reiken, S. and Marks, A. (2004) Circ Res 94, e61–e70.

242. Xiao, B., Jiang, M., Zhao, M., Yang, D., Sutherland, C., Lai, F., Walsh, M., Warltier, D., Cheng,

H. and Chen, S. (2005) Circ Res 96, 847–855.

243. Reiken, S., Lacampagne, A., Zhou, H., Kherani, A., Lehnart, S., Ward, C., Huang, F.,

Gaburjakova, M., Gaburjakova, J., Rosemblit, N., Warren, M., He, K., Yi, G., Wang, J.,

Burkhoff, D., Vassort, G. and Marks, A. (2003) J Cell Biol 160, 919–928.

244. Xiao, B., Zhong, G., Obayashi, M., Yang, D., Chen, K., Walsh, M., Shimoni, Y., Cheng, H., ter

Keurs, H. and Chen, S. (2006) Biochem J 396, 7–16.

245. Lehnart, S., Wehrens, X., Reiken, S., Warrier, S., Belevych, A., Harvey, R., Richter, W., Jin, S.,

Conti, M. and Marks, A. (2005) Cell 123, 25–35.

246. Wehrens, X., Lehnart, S., Reiken, S., Vest, J., Wronska, A. andMarks, A. (2006) Proc Natl Acad Sci

USA 103, 511–518.

247. Stange, M., Xu, L., Balshaw, D., Yamaguchi, N. and Meissner, G. (2003) J Biol Chem 278,

51693–51702.

248. Carter, S., Colyer, J. and Sitsapesan, R. (2006) Circ Res 98, 1506–1513.

249. Valdivia, H., Kaplan, J., Ellis-Davies, G. and Lederer, W. (1995) Science 267, 1997–2000.

250. Mayrleitner, M., Chandler, R., Schindler, H. and Fleischer, S. (1995) Cell Calcium 18, 197–206.

251. Patel, J., Coronado, R. and Moss, R. (1995) Circ Res 77, 943–949.

252. Uehara, A., Yasukochi, M., Mejia-Alvarez, R., Fill, M. and Imanaga, I. (2002) Pflugers Arch 444,

202–212.

253. Lokuta, A., Rogers, T., Lederer, W. and Valdivia, H. (1995) J Physiol 487, 609–622.

254. Terentyev, D., Viatchenko-Karpinski, S., Gyorke, I., Terentyeva, R. and Gyorke, S. (2003)

J Physiol 552, 109–118.

255. Bers, D. (2002) Nature 415, 198–205.

256. Lindegger, N. and Niggli, E. (2005) J Physiol 565, 801–813.

257. Ginsburg, K. and Bers, D. (2004) J Physiol 556, 463–480.

258. Li, Y., Kranias, E., Mignary, G. and Bers, D. (2002) Circ Res 90, 309–316.

259. Guo, T., Zhang, T., Mestril, R. and Bers, D. (2006) Circ Res. 99, 398–406.

260. Li, L., Satoh, H., Ginsburg, K. and Bers, D. (1997) J Physiol 501, 17–31.

261. Maier, L., Zhang, T., Chen, L., DeSantiago, J., Brown, J. and Bers, D. (2003) Circ Res 92, 904–911.


262. Kohlhaas, M., Zhang, T., Seidler, T., Zibrova, D., Dybkova, N., Steen, A., Wagner, S., Chen, L.,

Brown, J., Bers, D. and Maier, L. (2006) Circ Res 98, 235–244.

263. Wu, Y., Colbran, R. and Anderson, M. (2001) Proc Natl Acad Sci USA 98, 2877–2881.

264. duBell, W., Lederer, W. and Rogers, T. (1996) J Physiol 493, 793–800.

265. Takekura, H., Takeshima, H., Nishimura, S., Takahashi, M., Tanabe, T., Flockerzi, V., Hofmann,

F. and Franzini-Armstrong, C. (1995) J Muscle Res Cell Motil 16, 465–480.

266. Yin, C. and Lai, F. (2000) Nat Cell Biol 2, 669–671.

267. Yin, C., Blayney, L. and Lai, F. (2005) J Mol Biol 349, 538–546.

268. Blayney, L., Zissimopoulos, S., Ralph, E., Abbot, E., Matthews, L. and Lai, F. (2004) J Biol Chem

279, 14639–14648.

269. Niggli, E. (1999) Annu Rev Physiol 61, 311–335.

270. Gao, L., Tripathy, A., Lu, X. and Meissner, G. (1997) FEBS Lett 412, 223–226.

271. Stewart, R., Zissimopoulos, S. and Lai, F. (2003) Biochem J 376, 795–799.

272. Chen, S., Airey, J. and MacLennan, D. (1993) J Biol Chem 268, 22642–22649.

273. Masumiya, H., Wang, R., Zhang, J., Xiao, B. and Chen, S. (2003) J Biol Chem 278,

3786–3792.

274. Yamamoto, T., El-Hayek, R. and Ikemoto, N. (2000) J Biol Chem 275, 11618–11625.

275. Lamb, G., Posterino, G., Yamamoto, T. and Ikemoto, N. (2001) Am J Physiol 281, C207–C214.

276. Shtifman, A., Ward, C., Yamamoto, T., Wang, J., Olbinski, B., Valdivia, H., Ikemoto, N. and

Schneider, M. (2002) J Gen Physiol 119, 15–32.

277. Yamamoto, T. and Ikemoto, N. (2002) Biochemistry 41, 1492–1501.

278. Kobayashi, S., Yamamoto, T., Parness, J. and Ikemoto, N. (2004) Biochem J 380, 561–569.

279. Bannister, M. and Ikemoto, M. (2006) Biochem J 394, 145–152.

280. Yamamoto, T. and Ikemoto, N. (2002) Biochem Biophys Res Commun 291, 1102–1108.

281. Oda, T., Yano, M., Yamamoto, T., Tokuhisa, T., Okuda, S., Doi, M., Ohkusa, T., Ikeda, Y.,

Kobayashi, S., Ikemoto, N. and Matsuzaki, M. (2005) Circulation 111, 3400–3410.

282. Yang, Z., Ikemoto, N., Lamb, G. and Steele, D. (2006) Cardiovasc Res 70, 475–485.

283. Zorzato, F., Menegazzi, P., Treves, S. and Ronjat, M. (1996) J Biol Chem 271, 22759–22763.

284. El-Hayek, R., Saiki, Y., Yamamoto, T. and Ikemoto, N. (1999) J Biol Chem 274, 33341–33347.

285. Paul-Pletzer, K., Yamamoto, T., Bhat, M., Ma, J., Ikemoto, N., Jimenez, L., Morimoto, H.,

Williams, P. and Parness, J. (2002) J Biol Chem 277, 34918–34923.

286. Kobayashi, S., Bannister, M., Gangopadhyay, J., Hamada, T., Parness, J. and Ikemoto, N. (2005) J

Biol Chem 280, 6580–6587.

287. Paul-Pletzer, K., Yamamoto, T., Ikemoto, N., Jimenez, L., Morimoto, H., Williams, P., Ma, J. and

Parness, J. (2005) Biochem J 387, 905–909.

288. Murayama, T., Oba, T., Kobayashi, S., Ikemoto, N. and Ogawa, Y. (2005) Am J Physiol 288,

C1222–C1230.

289. Zhu, X., Ghanta, J., Walker, J., Allen, P. and Valdivia, H. (2004) Cell Calcium 35, 165–177.

290. Rodney, G., Wilson, G. and Schneider, M. (2005) J Biol Chem 280, 11713–11722.

291. Xiong, L., Zhang, J.-Z., He, R. and Hamilton, S. (2006) Biophys J 90, 173–182.

292. Gangopadhyay, J. and Ikemoto, N. (2006) Biophys J 90, 2015–2026.

293. Moore, C., Zhang, J.-Z. and Hamilton, S. (1999) J Biol Chem 274, 36831–36834.

294. Zhang, H., Zhang, J.-Z., Danila, C. and Hamilton, S. (2003) J Biol Chem 278, 8348–8355.

295. Treves, S., Scutari, E., Robert, M., Groh, S., Ottolia, M., Prestipino, G., Ronjat, M. and Zorzato, F.

(1997) Biochemistry 36, 11496–11503.

296. Nelson, T., Zhao, W., Yuan, S., Favit, A., Pozzo-Miller, L. and Alkon, D. (1999) Biochem J 341,

423–433.

297. Feng, W., Tu, J., Yang, T., Vernon, P., Allen, P., Worley, P. and Pessah, I. (2002) J Biol Chem 277,

44722–44730.

298. Zissimopoulos, S., West, D., Williams, A. and Lai, F. (2006) J Cell Sci 119, 2386–2397.


299. Tanabe, T., Takeshima, H., Mikami, A., Flockerzi, V., Takahashi, H., Kangawa, K., Kojima, M.,

Matsuo, H., Hirose, T. and Numa, S. (1987) Nature 328, 313–318.

300. Mikami, A., Imoto, K., Tanabe, T., Niidome, T., Mori, Y., Takeshima, H., Narumiya, S. and

Numa, S. (1989) Nature 340, 230–233.

301. Nakai, J., Tanabe, T., Konno, T., Adams, B. and Beam, K. (1998) J Biol Chem 273, 24983–24986.

302. Proenza, C., Wilkens, C. and Beam, K. (2000) J Biol Chem 275, 29935–29937.

303. Ahern, C., Bhattacharya, D., Mortenson, L. and Coronado, R. (2001) Biophys J 81, 3294–3307.

304. Wilkens, C., Kasielke, N., Flucher, B., Beam, K. and Grabner, M. (2001) Proc Natl Acad Sci USA

98, 5892–5897.

305. Kugler, G., Weiss, R., Flucher, B. and Grabner, M. (2004) J Biol Chem 279, 4721–4728.

306. Lu, X., Xu, L. and Meissner, G. (1994) J Biol Chem 269, 6511–6516.

307. O’Reilly, F., Robert, M., Jona, I., Szegedi, C., Albrieux, M., Geib, S., De Waard, M., Villaz, M. and

Ronjat, M. (2002) Biophys J 82, 145–155.

308. Lu, X., Xu, L. and Meissner, G. (1995) J Biol Chem 270, 18459–18464.

309. El-Hayek, R., Antoniu, B., Wang, J., Hamilton, S. and Ikemoto, N. (1995) J Biol Chem 270,

22116–22118.

310. Saiki, Y., El-Hayek, R. and Ikemoto, N. (1999) J Biol Chem 274, 7825–7832.

311. Lamb, G., El-Hayek, R., Ikemoto, N. and Stephenson, D. (2000) Am J Physiol 279, C891–C905.

312. Stange, M., Tripathy, A. and Meissner, G. (2001) Biophys J 81, 1419–1429.

313. Yamamoto, T., Rodriguez, J. and Ikemoto, N. (2002) J Biol Chem 277, 993–1001.

314. Haarmann, C., Green, D., Casarotto, M., Laver, D. and Dulhunty, A. (2003) Biochem J 372,

305–316.

315. Haarmann, C., Dulhunty, A. and Laver, D. (2005) Biochem J 387, 429–436.

316. El-Hayek, R. and Ikemoto, N. (1998) Biochemistry 37, 7015–7020.

317. Dulhunty, A., Laver, D., Gallant, E., Casarotto, M., Pace, S. and Curtis, S. (1999) Biophys J 77,

189–203.

318. Gurrola, G., Arevalo, C., Sreekumar, R., Lokuta, A., Walker, J. and Valdivia, H. (1999) J Biol

Chem 274, 7879–7886.

319. Casarotto, M., Green, D., Pace, S., Curtis, S. and Dulhunty, A. (2001) Biophys J 80, 2715–2726.

320. Slavik, K., Wang, J., Aghdasi, B., Zhang, J., Mandel, F., Malouf, N. and Hamilton, S. (1997) Am J

Physiol 272, C1475–C1481.

321. Leong, P. and MacLennan, D. (1998) J Biol Chem 273, 29958–29964.

322. Takekura, H., Paolini, C., Franzini-Armstrong, C., Kugler, G., Grabner, M. and Flucher, B. (2004)

Mol Biol Cell 15, 5408–5419.

323. Carbonneau, L., Bhattacharya, D., Sheridan, D. and Coronado, R. (2005) Biophys J 89, 243–255.

324. Cheng, W., Altafaj, X., Ronjat, M. and Coronado, R. (2005) Proc Natl Acad Sci USA 102,

19225–19230.

325. Nakai, J., Ogura, T., Protasi, F., Franzini-Armstrong, C., Allen, P. and Beam, K. (1997) Proc Natl

Acad Sci USA 94, 1019–1022.

326. Fessenden, J., Wang, Y., Moore, R., Chen, S., Allen, P. and Pessah, I. (2000) Biophys J 79,

2509–2525.

327. Leong, P. and MacLennan, D. (1998) J Biol Chem 273, 7791–7794.

328. Nakai, J., Sekiguchi, N., Rando, T., Allen, P. and Beam, K. (1998) J Biol Chem 273, 13403–13406.

329. Protasi, F., Paolini, C., Nakai, J., Beam, K., Franzini-Armstrong, C. and Allen, P. (2002) Biophys J

83, 3230–3244.

330. Perez, C., Voss, A., Pessah, I. and Allen, P. (2003) Biophys J 84, 2655–2663.

331. Perez, C., Mukherjee, S. and Allen, P. (2003) J Biol Chem 278, 39644–39652.

332. Yamazawa, T., Takeshima, H., Shimuta, M. and Iino, M. (1997) J Biol Chem 272, 8161–8164.

333. Altafaj, X., Cheng, W., Esteve, E., Urbani, J., Grunwald, D., Sabatier, J., Coronado, R., De Waard,

M. and Ronjat, M. (2005) J Biol Chem 280, 4013–4016.


334. Proenza, C., O’Brien, J., Nakai, J., Mukherjee, S., Allen, P. and Beam, K. (2002) J Biol Chem 277,

6530–6535.

335. Sencer, S., Papineni, R., Halling, D., Pate, P., Krol, J., Zhang, J. and Hamilton, S. (2001) J Biol

Chem 276, 38237–38241.

336. Kay, J. (1996) Biochem J 314, 361–385.

337. Michnick, S., Rosen, M., Wandless, T., Karplus, M. and Schreiber, S. (1991) Science 252, 836–839.

338. Van Duyne, G., Standaert, R., Schreiber, S. and Clardy, J. (1991) J Am Chem Soc 113, 7433–7434.

339. Deivanayagam, C., Carson, M., Thotakura, A., Narayama, S. and Chodavarapu, R. (2000) Acta

Crystallogr D 56, 266–271.

340. Lam, E., Martin, M., Timerman, A., Sabers, C., Fleischer, S., Lukas, T., Abraham, R., O’Keefe, S.,

O’Neill, E. and Wiederrecht, G. (1995) J Biol Chem 270, 26511–26522.

341. Jayaraman, T., Brillantes, A., Timerman, A., Fleischer, S., Erdjument-Bromage, H., Tempst, P. and

Marks, A. (1992) J Biol Chem 267, 9474–9477.

342. Timerman, A., Wiederrecht, G., Marcy, A. and Fleischer, S. (1995) J Biol Chem 270, 2451–2459.

343. Timerman, A., Onoue, H., Xin, H., Barg, S., Copello, J., Wiederrecht, G. and Fleischer, S. (1996)

J Biol Chem 271, 20385–20391.

344. Qi, Y., Ogunbunmi, E., Freund, E., Timerman, A. and Fleischer, S. (1998) J Biol Chem 273,

34813–34819.

345. Lee, E., Rho, S., Kwon, S., Eom, S., Allen, P. and Kim, D. (2004) J Biol Chem 279, 26481–26488.

346. Jones, J., Reynolds, D., Lai, F. and Blayney, L. (2005) J Cell Sci 118, 4613–4619.

347. Mackrill, J., O’Driscoll, S., Lai, F. and McCarthy, T. (2001) Biochem Biophys Res Commun 285,

52–57.

348. Timerman, A., Jayaraman, T., Wiederrecht, G., Onoue, H., Marks, A. and Fleischer, S. (1994)

Biochem Biophys Res Commun 198, 701–706.

349. Jeyakumar, L., Ballester, L., Cheng, D., McIntyre, J., Chang, P., Olivey, H., Rollins-Smith, L.,

Barnett, J., Murray, K., Xin, H.-B. and Fleischer, S. (2001) Biochem Biophys Res Commun 281,

979–986.

350. Xin, H., Rogers, K., Qi, Y., Kanematsu, T. and Fleischer, S. (1999) J Biol Chem 274, 15315–15319.

351. Huang, F., Shan, J., Reiken, S., Wehrens, X. and Marks, A. (2006) Proc Natl Acad Sci USA 103,

3456–3461.

352. Sharma, M., Jeyakumar, L., Fleischer, S. and Wagenknecht, T. (2006) Biophys J 90, 164–172.

353. Bultynck, G., Rossi, D., Callewaert, G., Missiaen, L., Sorrentino, V., Parys, J. and De Smet, H.

(2001) J Biol Chem 276, 47715–47724.

354. Timerman, A., Ogunbumni, E., Freund, E., Wiederrecht, G., Marks, A. and Fleischer, S. (1993)

J Biol Chem 268, 22992–22999.

355. Kaftan, E., Marks, A. and Ehrlich, B. (1996) Circ Res 78, 990–997.

356. Ahern, G., Junankar, P. and Dulhunty, A. (1997) Biophys J 72, 146–162.

357. Cameron, A., Nucifora, F., Fung, E., Livingston, D., Aldape, R., Ross, C. and Snyder, S. (1997)

J Biol Chem 272, 27582–27588.

358. Bultynck, G., De Smet, P., Rossi, D., Callewaert, G., Missiaen, L., Sorrentino, V., De Smet, H. and

Parys, J. (2001) Biochem J 354, 413–422.

359. Gaburjakova, M., Gaburjakova, J., Reiken, S., Huang, F., Marx, S., Rosemblit, N. and Marks, A.

(2001) J Biol Chem 276, 16931–16935.

360. Avila, G., Lee, E., Perez, C., Allen, P. and Dirksen, R. (2003) J Biol Chem 278, 22600–22608.

361. Van Acker, K., Bultynck, G., Rossi, D., Sorrentino, V., Boens, N., Missiaen, L., De Smedt, H.,

Parys, J. and Callewaert, G. (2004) J Cell Sci 117, 1129–1137.

362. Zissimopoulos, S. and Lai, F. (2005) Cell Biochem Biophys 43, 203–220.

363. Xiao, B., Sutherland, C., Walsh, M. and Chen, S. (2004) Circ Res 94, 487–495.

364. Zissimopoulos, S. and Lai, F. (2005) J Biol Chem 280, 5475–5485.

365. Samso, M., Shen, X. and Allen, P. (2006) J Mol Biol 356, 917–927.


366. Ahern, G., Junankar, P. and Dulhunty, A. (1994) FEBS Lett 352, 369–374.

367. Brillantes, A., Ondrias, K., Scott, A., Kobrinsky, E., Ondriasova, E., Moschella, M., Jayaraman, T.,

Landers, M., Ehrlich, B. and Marks, A. (1994) Cell 77, 513–523.

368. Mayrleitner, M., Timerman, A., Wiederrecht, G. and Fleischer, S. (1994) Cell Calcium 15, 99–108.

369. Barg, S., Copello, J. and Fleisher, S. (1997) Am J Physiol 272, C1726–C1733.

370. Shou, W., Aghdasi, B., Armstrong, D., Guo, Q., Bao, S., Charng, M., Mathews, L., Schneider, M.,

Hamilton, S. and Matzuk, M. (1998) Nature 391, 489–492.

371. Xiao, R., Valdivia, H., Bogdanov, K., Valdivia, C., Lakatta, E. and Cheng, H. (1997) J Physiol 500,

343–354.

372. Wehrens,X., Lenhart, S.,Huang, F., Vest, J., Reiken, S.,Mohler, P., Sun, J.,Guatimosim, S., Song, L.-S.,

Rosemblit, N., D’Armiento, J., Napolitano, C.,Memmi,M., Priori, S., Lederer,W. andMarks, A. (2003)

Cell 113, 829–840.

373. Marx, S., Ondrias, K. and Marks, A. (1998) Science 281, 818–821.

374. Marx, S., Gaburjakova, J., Gaburjakova, M., Henrikson, C., Ondrias, K. andMarks, A. (2001) Circ

Res 88, 1151–1158.

375. Brooksbank, R., Badenhorst, M., Isaacs, H. and Savage, N. (1998) Anesthesiology 89, 693–698.

376. Tang, W., Ingalls, C., Durham, W., Snider, J., Reid, M., Wu, G., Matzuk, M. and Hamilton, S.

(2004) FASEB J 18, 1597–1599.

377. Lamb, G. and Stephenson, D. (1996) J Physiol 494, 569–576.

378. Avila, G. and Dirksen, R. (2005) Cell Calcium 38, 35–44.

379. McCall, E., Li, L., Satoh, H., Shannon, T., Blatter, L. and Bers, D. (1996) Circ Res 79, 1110–1121.

380. Xin, H.-B., Senbonmatsu, T., Cheng, D.-S., Wang, Y.-X., Copello, J., Ji, G.-J., Collier, M., Deng,

K.-Y., Jeyakumar, L., Magnuson, M., Inagami, T., Kotlikoff, M. and Fleischer, S. (2002) Nature

416, 334–337.

381. George, C., Sorathia, R., Bertrand, B. and Lai, F. (2003) Biochem J 370, 579–589.

382. Goonasekera, S., Chen, S. and Dirksen, R. (2005) Am J Physiol 289, C1476–C1484.

383. Prestle, J., Janssen, P., Janssen, A., Zeitz, O., Lehnart, S., Bruce, L., Smith, G. and Hasenfuss, G.

(2001) Circ Res 88, 188–194.

384. Loughrey, C., Seidler, T., Miller, S., Prestle, J., MacEachern, K., Reynolds, D., Hasenfuss, G. and

Smith, G. (2004) J Physiol 556, 919–934.

385. Gomez, A., Schuster, I., Fauconnier, J., Prestle, J., Hasenfuss, G. and Richard, S. (2004) Am J

Physiol 287, H1987–H1993.

386. Chin, D. and Means, A. (2000) Trends Cell Biol 10, 322–328.

387. Moore, C., Rodney, G., Zhang, J.-Z., Santacruz-Toloza, L., Strasburg, G. and Hamilton, S. (1999)

Biochemistry 38, 8532–8537.

388. Rodney, G., Williams, B., Strasburg, G., Beckingham, K. and Hamilton, S. (2000) Biochemistry 39,

7807–7812.

389. Fruen, B., Bardy, J., Byrem, T., Strasburg, G. and Louis, C. (2000) Am J Physiol 279, C724–C733.

390. Balshaw, D., Xu, L., Yamaguchi, N., Pasek, D. and Meissner, G. (2001) J Biol Chem 276,

20144–20153.

391. Yamaguchi, N., Xu, L., Pasek, D., Evans, K., Chen, S. and Meissner, G. (2005) Biochemistry 44,

15074–15081.

392. Yang, H., Reedy, M., Burke, C. and Strasburg, G. (1994) Biochemistry 33, 518–525.

393. Tripathy, A., Xu, L., Mann, G. and Meissner, G. (1995) Biophys J 69, 106–119.

394. Chen, S. and MacLennan, D. (1994) J Biol Chem 269, 22698–22704.

395. Menegazzi, P., Larini, F., Treves, S., Guerrini, R., Quadroni, M. and Zorzato, F. (1994)


396. Rodney, G., Moore, C., Williams, B., Zhang, J., Krol, J., Pedersen, S. and Hamilton, S. (2001) J Biol

Chem 276, 2069–2074.

397. Yamaguchi, N., Xin, C. and Meissner, G. (2001) J Biol Chem 276, 22579–22585.


398. Xiong, L., Newman, R., Rodney, G., Thomas, O., Zhang, J., Persechini, A., Shea, M. and

Hamilton, S. (2002) J Biol Chem 277, 40862–40870.

399. Yamaguchi, N., Xu, L., Pasek, D., Evans, K. and Meissner, G. (2003) J Biol Chem 278,

23480–23486.

400. Yamaguchi, N., Xu, L., Evans, K., Pasek, D. and Meissner, G. (2004) J Biol Chem 279, 36433–36439.

401. Meissner, G. (1986) Biochemistry 25, 244–251.

402. Smith, J., Rousseau, E. and Meissner, G. (1989) Circ Res 64, 352–359.

403. Fuentes, O., Valdivia, C., Vaughan, D., Coronado, R. and Valdivia, H. (1994) Cell Calcium 15,

305–316.

404. Fruen, B., Black, D., Bloomquist, R., Bardy, J., Johnson, J., Louis, C. and Balog, E. (2003)


405. O’Connell, K., Yamaguchi, N., Meissner, G. and Dirksen, R. (2002) J Gen Physiol 120, 337–347.

406. Ikemoto, T., Takeshima, H., Iino, M. and Endo, M. (1998) Pflugers Arch 437, 43–48.

407. Xu, L. and Meissner, G. (2004) Biophys J 86, 797–804.

408. Rodney, G. and Schneider, M. (2003) Biophys J 85, 921–932.

409. Anderson, M. (2002) J Cardiovasc Electrophysiol 13, 195–197.

410. Maki, M., Kitaura, Y., Satoh, H., Ohkouchi, S. and Shibata, H. (2002) Biochim Biophys Acta 1600,

51–60.

411. Meyers, M., Pickel, V., Sheu, S., Sharma, V., Scotto, K. and Fishman, G. (1995) J Biol Chem 270,

26411–26418.

412. Meyers, M., Fischer, A., Sun, Y., Lopes, C., Rohacs, T., Nakamura, T., Zhou, Y., Lee, P.,

Altschuld, R., McCune, S., Coetzee, W. and Fishman, G. (2003) J Biol Chem 278, 28865–28871.

413. Farrell, E., Antaramian, A., Rueda, A., Gomez, A. and Valdivia, H. (2003) J Biol Chem 278,

34660–34666.

414. Meyers, M., Puri, T., Chien, A., Gao, T., Hus, P.-H., Hosey, M. and Fishman, G. (1998) J Biol

Chem 273, 18930–18935.

415. Lokuta, A., Meyers, M., Sander, P., Fishman, G. and Valdivia, H. (1997) J Biol Chem 272,

25333–25338.

416. Seidler, T., Miller, S., Loughrey, C., Kania, A., Burow, A., Kettlewell, S., Teucher, N., Wagner, S.,

Kogler, H., Meyers, M., Hasenfuss, G. and Smith, G. (2003) Circ Res 93, 132–139.

417. Suarez, J., Belke, D., Gloss, B., Dieterle, T., McDonough, P., Kim, Y., Brunton, L. andDillmann, D.

(2004) Am J Physiol 286, H68-H75.

418. Frank, K., Block, B., Ding, Z., Krause, D., Hattebuhr, N., Malik, A., Brixius, K., Hajjar, R.,

Schrader, J. and Schwinger, R. (2005) J Mol Cell Cardiol 38, 607–615.

419. Matsumoto, T., Hisamatsu, Y., Ohkusa, T., Inoue, N., Sato, T., Suzuki, S., Ikeda, Y. and

Matsuzaki, M. (2005) Basic Res Cardiol 100, 250–262.

420. Ilari, A., Johnson, K., Nastopoulos, V., Verzili, D., Zamparelli, C., Colotti, G., Tsernoglou, D. and

Chiancone, E. (2002) J Mol Biol 317, 447–458.

421. MacLennan, D. and Wong, P. (1971) Proc Natl Acad Sci USA 68, 1231–1235.

422. Ikemoto, N., Bhatnagar, G., Nagy, B. and Gergely, J. (1972) J Biol Chem 247, 7835–7837.

423. Wang, S., Trumble, W., Liao, H., Wesson, C., Dunker, A. and Kang, C. (1998) Nat Struct Biol 5,

476–483.

424. Fliegel, L., Ohnishi, M., Carpenter, M., Khanna, V., Reithmeier, R. and MacLennan, D. (1987)

Proc Natl Acad Sci USA 84, 1167–1171.

425. Scott, B., Simmerman, H., Collins, J., Nadal-Ginard, B. and Jones, L. (1988) J Biol Chem 263,

8958–8964.

426. Fryer, M. and Stephenson, D. (1996) J Physiol 493, 357–370.

427. Park, H., Park, I., Kim, E., Youn, B., Fields, K., Dunker, A. and Kang, C. (2004) J Biol Chem 279,

18026–18033.

428. Guo, W. and Campbell, K. (1995) J Biol Chem 270, 9027–9030.


429. Zhang, L., Kelley, J., Schmeisser, G., Kobayashi, Y. and Jones, L. (1997) J Biol Chem 272,

23389–23397.

430. Shin, D., Ma, J. and Kim, D. (2000) FEBS Lett 486, 178–182.

431. Beard, N., Sakowska, M., Dulhunty, A. and Laver, D. (2002) Biophys J 82, 310–320.

432. Herzog, A., Szegedi, C., Jona, I., Herberg, F. and Varsanyi, M. (2000) FEBS Lett 472,

73–77.

433. Brandt, N., Caswell, A., Wen, S. and Talvenheimo, J. (1990) J Membr Biol 113, 237–251.

434. Knudson, C., Stang, K., Moomaw, C., Slaughter, C. and Campbell, K. (1993) J Biol Chem 268,

12646–12654.

435. Marty, I., Thevenon, D., Scotto, C., Groh, S., Sainnier, S., Robert, M., Grunwald, D. and Villaz, M.

(2000) J Biol Chem 275, 8206–8212.

436. Vassilopoulos, S., Thevenon, D., Rezgui, S., Brocard, J., Chapel, A., Lacampagne, A., Lunardi, J.,

De Waard, M. and Marty, I. (2005) J Biol Chem 280, 28601–28609.

437. Guo, W., Jorgensen, A., Jones, L. and Campbell, K. (1996) J Biol Chem 271, 458–465.

438. Kobayashi, Y. and Jones, L. (1999) J Biol Chem 274, 28660–28668.

439. Marty, I., Robert, M., Ronjat, M., Bally, I., Arlaud, G. and Villaz, M. (1995) Biochem J 307,

769–774.

440. Caswell, A., Brandt, N., Brunschwig, J. and Purkerson, S. (1991) Biochemistry 30, 7507–7513.

441. Groh, S., Marty, I., Ottolia, M., Prestipino, G., Chapel, A., Villaz, M. and Ronjat, M. (1999) J Biol

Chem 274, 12278–12283.

442. Lee, J., Rho, S.-H., Shin, D., Cho, C., Park, W., Eom, S., Ma, J. and Kim, D. (2004) J Biol Chem

279, 6994–7000.

443. Kobayashi, Y., Alseikhan, B. and Jones, L. (2000) J Biol Chem 275, 17639–17646.

444. Jones, L., Zhang, L., Sanborn, K., Jorgensen, A. and Kelley, J. (1995) J Biol Chem 270,

30787–30796.

445. Ohkura, M., Furukawa, K., Fujimori, H., Kuruma, A., Kawano, S., Hiraoka, M., Kuniyasu, A.,

Nakayama, H. and Ohizumi, Y. (1998) Biochemistry 37, 12987–12993.

446. Kirchhefer, U., Neumann, J., Baba, H., Begrow, F., Kobayashi, Y., Reinke, U., Schmitz, W. and

Jones, L. (2001) J Biol Chem 276, 4142–4149.

447. Kirchhefer, U., Jones, L., Begrow, F., Boknik, P., Hein, L., Lohse, M., Riemann, B., Schmitz, W.,

Stypmann, J. and Neumann, J. (2004) Cardiovasc Res 62, 122–134.

448. Hong, C., Cho, M., Kwak, Y., Song, C., Lee, Y., Lim, J., Kwon, Y., Chae, S. and Kim, D. (2002)

FASEB J 16, 1310–1312.

449. Kirchhefer, U., Neumann, J., Bers, D., Buchwalow, I., Fabritz, L., Hanske, G., Justus, I., Riemann,

B., Schmitz, W. and Jones, L. (2003) Cardiovasc Res 59, 369–379.

450. Terentyev, D., Cala, S., Houle, T., Viatchenko-Karpinski, S., Gyorke, I., Terentyeva, R., Williams,

S. and Gyorke, S. (2005) Circ Res 96, 651–658.

451. Rezgui, S., Vassilopoulos, S., Brocard, J., Platel, J., Bouron, A., Arnoult, C., Oddoux, S., Garcia,

L., De Waard, M. and Marty, I. (2005) J Biol Chem 280, 39302–39308.

452. Kawasaki, T. and Kasai, M. (1994) Biochem Biophys Res Commun 199, 1120–1127.

453. Szegedi, C., Sarkozi, S., Herzog, A., Jona, I. and Varsanyi, M. (1999) Biochem J 337, 19–22.

454. Jones, L., Suzuki, Y., Wang, W., Kobayashi, Y., Ramesh, V., Franzini-Armstrong, C., Cleemann,

L. and Morad, M. (1998) J Clin Invest 101, 1385–1393.

455. Sato, Y., Ferguson, D., Sako, H., Dorn, G., Kadambi, V., Yatani, A., Hoit, B., Walsh, R. and

Kranias, E. (1998) J Biol Chem 273, 28470–28477.

456. Wang, W., Cleemann, L., Jones, L. and Morad, M. (2000) J Physiol 524, 399–414.

457. Miller, S., Currie, S., Loughrey, C., Kettlewell, S., Seidler, T., Reynolds, D., Hasenfuss, G. and

Smith, G. (2005) Cardiovasc Res 67, 667–677.

458. Terentyev, D., Viatchenko-Karpinski, S., Gyorke, I., Volpe, P., Williams, S. and Gyorke, S. (2003)

Proc Natl Acad Sci USA 100, 11759–11764.


459. Shin, D., Pan, Z., Kim, E., Lee, J., Bhat, M., Parness, J., Kim, D. andMa, J. (2003) J Biol Chem 278,

3286–3292.

460. Viatchenko-Karpinski, S., Terentyev, D., Gyorke, I., Terentyeva, R., Volpe, P., Priori, S.,

Napolitano, C., Nori, A., Williams, S. and Gyorke, S. (2004) Circ Res 94, 471–477.

461. Terentyev, D., Nori, A., Santoro, M., Viatchenko-Karpinski, S., Kubalova, Z., Gyorke, I.,

Terentyeva, R., Vedamoorthyrao, S., Blom, N., Valle, G., Napolitano, C., Williams, S., Volpe, P.,

Priori, S. and Gyorke, S. (2006) Circ Res 98, 1151–1158.

462. Houle, T., Ram, M. and Cala, S. (2004) Cardiovasc Res 64, 227–233.

463. Liew, C.-C. and Dzau, V. (2004) Nat Rev Genet 5, 811–825.

464. Bers, D., Eisner, D. and Valdivia, H. (2003) Circ Res 93, 487–490.

465. Shannon, T., Pogwizd, S. and Bers, D. (2003) Circ Res 93, 592–594.

466. Kubalova, Z., Terentyev, D., Viatchenko-Karpinski, S., Nishijima, Y., Gyorke, I., Terentyeva, R.,

da Cunha, D., Sridhar, A., Feldman, D., Hamlin, R., Carnes, C. and Gyorke, S. (2005) Proc Natl

Acad Sci USA 102, 14104–14109.

467. Reiken, S., Gaburjakova, M., Gaburjakova, J., He, K., Prieto, A., Becker, E., Yi, G., Wang, J.,

Burkhoff, D. and Marks, A. (2001) Circulation 104, 2843–2848.

468. Reiken, S., Gaburjakova, M., Guatimosim, S., Gomez, A., D’Armiento, J., Burkhoff, D., Wang, J.,

Vassort, G., Lederer, W. and Marks, A. (2003) J Biol Chem 278, 444–453.

469. Reiken, S., Wehrens, X., Vest, J., Barbone, A., Klotz, S., Mancini, D., Burkhoff, D. and Marks, A.

(2003) Circulation 107, 2459–2466.

470. Wehrens, X., Lehnart, S., Reiken, S., Deng, S., Vest, J., Cervantes, D., Coromilas, J., Landry, D.

and Marks, A. (2004) Science 304, 292–296.

471. Wehrens, X., Lehnart, S., Reiken, S., Van der Nagel, R., Morales, R., Sun, J., Cheng, Z., Deng, S.,

De Windt, L., Landry, D. and Marks, A. (2005) Proc Natl Acad Sci USA 102, 9607–9612.

472. Yano, M., Ono, K., Ohkusa, T., Suetsugu, M., Kohno, M., Hisaoka, T., Kobayashi, S., Hisamatsu,

Y., Yamamoto, T., Kohno, M., Noguchi, N., Takasawa, S., Okamoto, H. andMatsuzaki, M. (2000)

Circulation 102, 2131–2136.

473. Ono, K., Yano, M., Ohkusa, T., Kohno, M., Hisaoka, T., Tanigawa, T., Kobayashi, S., Kohno, M.

and Matsuzaki, M. (2000) Cardiovasc Res 48, 323–331.

474. Doi, M., Yano, M., Kobayashi, S., Kohno, M., Tokuhisa, T., Okuda, S., Suetsugu, M., Hisamatsu,

Y., Ohkusa, T., Kohno, M. and Matsuzaki, M. (2002) Circulation 105, 1374–1379.

475. Yano, M., Kobayashi, S., Kohno, M., Doi, M., Tokuhisa, T., Okuda, S., Suetsugu, M.,

Hisaoka, T., Obayashi, M., Ohkusa, T., Kohno, M. and Matsuzaki, M. (2003) Circulation

107, 477–484.

476. Jiang, M., Lokuta, A., Farrell, E., Wolff, M., Haworth, R. and Valdivia, H. (2002) Circ Res 91,

1015–1022.

477. Obayashi, M., Xiao, B., Stuyvers, B., Davidoff, A., Mei, J., Chen, S. and ter Keurs, H. (2006)


478. Ai, X., Curran, J., Shannon, T., Bers, D. and Pogwizd, S. (2005) Circ Res 97, 1314–1322.

479. Zhang, T., Maier, L., Dalton, N., Miyamoto, S., Ross, J.J., Bers, D. and Brown, J. (2003) Circ Res

92, 912–919.

480. Wang, W., Zhu, W., Wang, S., Yang, D., Crow, M., Xiao, R.-P. and Cheng, H. (2004) Circ Res 95,

798–806.

481. Zhang, R., Khoo, M., Wu, Y., Yang, Y., Grueter, C., Ni, G., Price, E.J., Thiel, W., Guatimosim, S.,

Song, L.-S., Madu, E., Shah, A., Vishnivetskaya, T., Atkinson, J., Gurevich, V., Salama, G.,

Lederer, W., Colbran, R. and Anderson, M. (2005) Nat Med 11, 409–417.

482. Francis, J., Sankar, V., Nair, V. and Priori, S. (2005) Heart Rhythm 2, 550–554.

483. Kontula, K., Laitinen, P., Lehtonen, A., Toivonen, L., Viitasalo, M. and Swan, H. (2005)


484. Jiang, D., Xiao, B., Zhang, L. and Chen, S. (2002) Circ Res 91, 218–225.

485. George, C., Higgs, G. and Lai, F. (2003) Circ Res 93, 531–540.


486. Jiang, D., Xiao, B., Yang, D., Wang, R., Choi, P., Zhang, L., Cheng, H. and Chen, S. (2004) Proc

Natl Acad Sci USA 101, 13062–13067.

487. Thomas, L., George, C. and Lai, F. (2004) Cardiovasc Res 64, 52–60.

488. Lehnart, S., Wehrens, X., Laitinen, P., Reiken, S., Deng, S., Cheng, Z., Landry, D., Kontula, K.,

Swan, H. and Marks, A. (2004) Circulation 109, 3208–3214.

489. Thomas, L., Lai, F. and George, C. (2005) Biochem Biophys Res Commun 331, 231–238.

490. Jiang, D., Wang, R., Xiao, B., Kong, H., Hunt, D., Choi, P., Zhang, L. and Chen, S. (2005) Circ Res

97, 1173–1181.

491. George, C., Jundi, H., Walters, N., Thomas, N., West, R. and Lai, F. (2006) Circ Res 98, 88–97.

492. Lehnart, S., Terrenoire, C., Reiken, S., Wehrens, X., Song, L., Tillman, E., Mancarella, S.,

Coromillas, J., Lederer, W., Kass, R. and Marks, A. (2006) Proc Natl Acad Sci USA 103,

7906–7910.

493. Liu, N., Colombi, B., Memmi, M., Zissimopoulos, S., Rizzi, N., Negri, S., Imbriani, M.,

Napolitano, C., Lai, F. and Priori, S. (2006) Circ Res 99, 292–298.

494. Cerrone, M., Colombi, B., Santoro, M., Raffaele di Barletta, M., Scelsi, M., Villani, L., Napolitano,

C. and Priori, S. (2005) Circ Res 96, e77–e82.

495. Kannankeril, P., Mitchell, B., Goonasekera, S., Chelu, M., Zhang, W., Sood, S., Kearney, D.,

Danila, C., De Biasi, M., Wehrens, X., Pautler, R., Roden, D., Taffet, G., Dirksen, R., Anderson,

M. and Hamilton, S. (2006) Proc Natl Acad Sci USA 103, 12179–12184.

496. Janse, M. (2004) Cardiovasc Res 61, 208–217.

497. Pogwizd, S. and Bers, D. (2004) Trends Cardiovasc Med 14, 61–66.

498. Treves, S., Anderson, A., Ducreux, S., Divet, A., Bleunven, C., Grasso, C., Paesante, S. and

Zorzato, F. (2005) Neuromuscul Disord 15, 577–587.

499. Gallant, E. and Lentz, L. (1992) Am J Physiol 262, C422–C426.

500. Otsu, K., Nishida, K., Kimura, Y., Kuzuya, T., Hori, M., Kamada, T. and Tada, M. (1994) J Biol

Chem 269, 9413–9415.

501. Treves, S., Larini, F., Menegazzi, P., Steinberg, T., Koval, M., Vilsen, B., Andersen, J. and Zorzato,

F. (1994) Biochem J 301, 661–665.

502. Owen, V., Taske, N. and Lamb, G. (1997) Am J Physiol 272, C203–C211.

503. Laver, D., Owen, V., Junankar, P., Taske, N., Dulhunty, A. and Lamb, G. (1997) Biophys J 73,

1913–1924.

504. Tong, J., Oyamada, H., Demaurex, N., Grinstein, S., McCarthy, T. and MacLennan, D. (1997) J

Biol Chem 272, 26332–26339.

505. Censier, K., Urwyler, A., Zorzato, F. and Treves, S. (1998) J Clin Invest 101, 1233–1242.

506. Dietze, B., Henke, J., Eichinger, H., Lehmann-Horn, F. and Melzer, W. (2000) J Physiol 526,

507–514.

507. Avila, G. and Dirksen, G. (2001) J Gen Physiol 118, 277–290.

508. Yang, T., Ta, T., Pessah, I. and Allen, P. (2003) J Biol Chem 278, 25722–25730.

509. Wehner, M., Rueffert, H., Koenig, F. and Olthoff, D. (2004) Neuromuscul Disord 14, 429–437.

510. Chelu, M., Goonasekera, S., Durham, W., Tang, W., Lueck, J., Riehl, J., Pessah, I., Zhang, P.,

Bhattacharjee, M., Dirksen, R. and Hamilton, S. (2005) FASEB J 20, 329–330.

511. Tong, J., McCarthy, T. and MacLennan, D. (1999) J Biol Chem 274, 693–702.

512. Tilgen, N., Zorzarto, F., Halliger-Keller, B., Muntoni, F., Sewry, C., Palmucci, L., Schneider, C.,

Hauser, E., Lehmann-Horn, F., Muller, C. and Treves, S. (2001) Hum Mol Genet 10, 2879–2887.

513. Avila, G., O’Connell, K. and Dirksen, R. (2003) J Gen Physiol 121, 277–286.

514. Zorzato, F., Yamaguchi, N., Xu, L., Meissner, G., Muller, C., Pouliquin, P., Muntoni, F., Sewry,

C., Girard, T. and Treves, S. (2003) Hum Mol Genet 12, 379–388.

515. Dirksen, R. and Avila, G. (2004) Biophys J 87, 3193–3204.

516. Brini, M., Manni, S., Pierobon, N., Du, G., Sharma, P., MacLennan, D. and Carafoli, E. (2005) J

Biol Chem 280, 15380–15389.


517. Lynch, P., Tong, J., Lehane, M., Mallet, A., Giblin, L., Heffron, J., Vaughan, P., Zafra, G.,

MacLennan, D. and McCarthy, T. (1999) Proc Natl Acad Sci USA 96, 4164–4169.

518. Ducreux, S., Zorzato, F., Muller, C., Sewry, C., Muntoni, F., Quinlivan, R., Restagno, G., Girard,

T. and Treves, S. (2004) J Biol Chem 279, 43838–43846.

519. Avila, G., O’Brien, J. and Dirksen, R. (2001) Proc Natl Acad Sci USA 98, 4215–4220.

520. Du, G., Khanna, V., Guo, X. and MacLennan, D. (2004) Biochem J 382, 557–564.


[new comprehensive biochemistry] calcium - a matter of life or death volume 41 || ryanodine receptor...

Documents